Methods for the high-resolution identification of solvent-accessible amide hydrogens in protein binding sites

ABSTRACT

The present invention provides methods whereby the positions of peptide amide groups that are labeled with a heavy hydrogen in a polypeptide or protein can be localized at high resolution. The methods are useful for determining which peptide amide groups in a polypeptide or protein are accessible to solvent, mapping the binding site and/or binding surface of a binding protein, and/or studying allosteric or other conformational changes in a polypeptide or protein which alter the rates at which certain peptide amide hydrogens exchange with solvent.

This application is a divisional of application Ser. No. 08/919,187,filed Aug. 19, 1997, now U.S. Pat. No. 6,291,189, which is acontinuation-in-part of application Ser. No. 08/895,330, entitled“Methods for Characterization of the Fine Structure of Protein BindingSites”, filed Jul. 16, 1997, now U.S. Pat. No. 6,331,400, which is acontinuation of application Ser. No. 08/240,593, filed May 10, 1994 nowU.S. Pat. No. 5,658,739, each of which is hereby incorporated herein inits entirety by reference.

BACKGROUND OF THE INVENTION

1. Field of the Invention

The present invention relates to the identification ofsolvent-accessible amide hydrogens in polypeptides or proteins. Themethods of the invention can be used to characterize the binding siteinvolved in binding between a binding protein and a binding partner, andto study other changes in a polypeptide or protein which alter the ratesat which hydrogen atoms exchange with solvent hydrogens, such as foldingphenomena and other structural changes.

2. Background Art

Limitations of Current Methods of Characterizing Protein Binding Sites

Considerable experimental work and time are required to preciselycharacterize a binding site. In general, the techniques which are theeasiest to use and which give the quickest answers, result in an inexactand only approximate idea of the nature of the critical structuralfeatures. Techniques in this category include the study ofproteolytically generated fragments of the protein which retain bindingfunction; recombinant DNA techniques, in which proteins are constructedwith altered amino acid sequence (site directed mutagenesis); epitopescanning peptide studies (construction of a large number of smallpeptides representing subregions of the intact protein followed by studyof the ability of the peptides to inhibit binding of the ligand toreceptor); covalent crosslinking of the protein to its binding partnerin the area of the binding site, followed by fragmentation of theprotein and identification of crosslinked fragments; and affinitylabeling of regions of the receptor which are located near the ligandbinding site of the receptor, followed by characterization of such“nearest neighbor” peptides. (Reviewed in 1, 2).

These techniques work best for the determination of the structure ofbinding subregions which are simple in nature, as when a single shortcontiguous stretch of polypeptide within a protein is responsible formost of the binding activity. However, for many protein-binding partnersystems of current interest, the structures responsible for binding onboth receptor and ligand or antibody are created by the complexinteraction of multiple non-contiguous peptide sequences. Thecomplexities of these interactions may confound conventional analyticaltechniques, as binding function is often lost as soon as one of the3-dimensional conformations of the several contributing polypeptidesequences is directly or indirectly perturbed.

The most definitive techniques for the characterization of the structureof receptor binding sites have been NMR spectroscopy and X-raycrystallography. While these techniques can ideally provide a precisecharacterization of the relevant structural features, they have majorlimitations, including inordinate amounts of time required for study,inability to study large proteins, and, for X-ray analysis, the need forprotein-binding partner crystals (Ref. 3).

Applicant's technology overcomes these limitations and allows the rapididentification of each of the specific polypeptides and amino acidswithin a protein which constitute its protein ligand binding site orantibody binding subregion in virtually any protein-ligand system orprotein antigen-antibody system, regardless of the complexity of thebinding sites present or the size of the proteins involved. Thistechnology is superior in speed and resolution to currently employedbiochemical techniques.

Hydrogen (Proton) Exchange

When a protein in its native folded state is incubated in bufferscontaining heavy hydrogen (tritium or deuterium) labeled water, heavyhydrogen in the buffer reversibly exchanges with normal hydrogen presentin the protein at acidic positions (for example, O—H, S—H, and N—Hgroups) with rates of exchange which are dependent on each exchangeablehydrogen's chemical environment, temperature, and most importantly, itsaccessibility to the tritiated water in the buffer. (Refs. 4, 5)Accessibility is determined in turn by both the surface(solvent-exposed) disposition of the hydrogen, and the degree to whichit is hydrogen-bonded to other regions of the folded protein. Simplystated, acidic hydrogen present on amino acid residues which are on theoutside (buffer-exposed) surface of the protein and which arehydrogen-bonded to solvent water will exchange more rapidly with heavyhydrogen in the buffer than will similar acidic hydrogen which areburied and hydrogen bonded within the folded protein.

Hydrogen exchange reactions can be greatly accelerated by both acid andbase-mediated catalysis, and the rate of exchange observed at anyparticular pH is the sum of both acid and base mediated mechanisms. Formany acidic hydrogen, a pH of 2.7 results in an overall minimum rate ofexchange (Ref. 6, pg. 238, FIGS. 3a-c, refs. 7-11). While hydrogens inprotein hydroxyl and amino groups exchange with tritium in buffer atmillisecond rates, the exchange rate of one particular acidic hydrogen,the peptide amide bond hydrogen, is considerably slower, having a halflife of exchange (when freely hydrogen bonded to solvent water) ofapproximately 0.5 seconds at 0° C., pH 7, which is greatly slowed to ahalf life of exchange of 70 minutes at 0° C. pH 2.7.

When peptide amide hydrogens are buried within a folded protein, or arehydrogen bonded to other parts of the protein, exchange half lives withsolvent hydrogens are often considerably lengthened, at times beingmeasured in hours to days. Hydrogen exchange at peptide amides is afully reversible reaction, and rates of on-exchange (solvent heavyhydrogen replacing protein-bound normal hydrogen) are identical to ratesof off-exchange (hydrogen replacing protein-bound heavy hydrogen) if thestate of a particular peptide amide within a protein, including itschemical environment and accessibility to solvent hydrogens, remainsidentical during on-exchange and off-exchange conditions.

Hydrogen exchange is commonly measured by performing studies withproteins and aqueous buffers that are differentially tagged with pairsof the three isotopic forms of hydrogen (¹H;Normal Hydrogen;²H;Deuterium; ³H;Tritium). If the pair of normal hydrogen and tritiumare employed, it is referred to as tritium exchange; if normal hydrogenand deuterium are employed, as deuterium exchange. Differentphysicochemical techniques are in general used to follow thedistribution of the two isotopes in deuterium versus tritium exchange.

Tritium Exchange Techniques

Tritium exchange techniques (where the amount of the isotope isdetermined by radioactivity measurements) have been extensively used forthe measurement of peptide amide exchange rates within an individualprotein (reviewed in 4). The rates of exchange of other acidic protons(OH, NH, SH) are so rapid that they cannot be followed in thesetechniques and all subsequent discussion refers exclusively to peptideamide proton exchange. In these studies, purified proteins areon-exchanged by incubation in buffers containing tritiated water forvarying periods of time, transferred to buffers free of tritium, and therate of off-exchange of tritium determined. By analysis of the rates oftritium on- and off-exchange, estimates of the numbers of peptide amideprotons in the protein whose exchange rates fall within particularexchange rate ranges can be made. These studies do not allow adetermination of the identity (location within the protein's primaryamino acid sequence) of the exchanging amide hydrogens measured.

Extensions of these techniques have been used to detect the presencewithin proteins of peptide amides which experienceallosterically-induced changes in their local chemical environment andto study pathways of protein folding (5, 12-14). For these studies,tritium on-exchanged proteins are allowed to off-exchange after theyhave experienced either an allosteric change in shape, or have undergonetime-dependent folding upon themselves, and the number of peptide amideswhich experience a change in their exchange rate subsequent to theallosteric/folding modifications determined. Changes in exchange rateindicate that alterations of the chemical environment of particularpeptide amides have occurred which are relevant to proton exchange(solvent accessibility, hydrogen bonding etc.). Peptide amides whichundergo an induced slowing in their exchange rate are referred to as“slowed amides” and if previously on-exchanged tritium is sufficientlyslowed in its off-exchange from such amides there results a “functionaltritium labeling” of these amides. From these measurements, inferencesare made as to the structural nature of the shape changes which occurredwithin the isolated protein. Again, determination of the identity of theparticular peptide amides experiencing changes in their environment isnot possible with these techniques.

Four groups of investigators have described technical extensions(collectively referred to as medium resolution tritium exchange) whichallow the locations of particular slowed, tritium labeled peptide amideswithin the primary sequence of small proteins to be localized to aparticular proteolytic fragment, though not to a particular amino acid.

Rosa and Richards were the first to describe and utilize mediumresolution tritium techniques in their studies of the folding ofribonuclease S protein fragments (15-17). However, the techniquesdescribed by Rosa and Richards were of marginal utility, primarily dueto their failure to optimize certain critical experimental steps(reviewed in 6, pg 238, 244). No studies employing related techniqueswere published until the work of Englander and co-workers in whichextensive modifications and optimizations of the Rosa and Richardstechnique were first described.

Englander's investigations utilizing tritium exchange have focusedexclusively on the study of allosteric changes which take place intetrameric hemoglobin (a subunit and b subunit 16 kD in size each) upondeoxygenation (6,18-21). In the Englander procedure, native hemoglobin(milligram quantities) in the oxygenated state is on-exchanged intritiated water of relatively low specific activity (2-100 mCi/ml). Thehemoglobin is then deoxygenated (inducing allosteric change),transferred to tritium-free buffers by gel permeation columnchromatography, and then allowed to out-exchange for 10-50 times theon-exchange time. On-exchanged tritium present on peptide amides whichexperience no change in exchange rate subsequent to the inducedallosteric change in hemoglobin structure off-exchanges at ratesidentical to its on-exchange rates, and therefore is almost totallyremoved from the protein after the long off-exchange period. However,peptide amides which experience slowing of their exchange ratesubsequent to the induced allosteric changes preferentially retain thetritium label during the period of off-exchange.

To localize (in terms of hemoglobin's primary sequence) the slowedamides bearing the residual tritium label, Englander thenproteolytically fragments the off-exchanged hemoglobin with the proteasepepsin, separates, isolates and identifies the various peptide fragmentsby reverse phase high pressure liquid chromatography (RP-HPLC), anddetermines which fragments bear the residual tritium label byscintillation counting. However, as the fragmentation of hemoglobinproceeds, each fragment's secondary and tertiary structure is lost andthe unfolded peptide amides become freely accessible to H₂O in thebuffer. At physiologic pH (>6), any amide-bound tritium label wouldleave the unfolded fragments within seconds. Englander thereforeperforms the fragmentation and HPLC peptide isolation procedures underconditions which he believes minimize peptide amide proton exchange,including cold temperature (4° C.) and use of phosphate buffers at pH2.7 (reviewed in 6). This technique has been used successfully byEnglander to coarsely identify and localize the peptidic regions ofhemoglobin α and β chains which participate in deoxygenation-inducedallosteric changes (18-21). The ability of the Englander technique tolocalize tritium labeled amides, while an important advance, remainslow; at the best, Englander reports that his technique localizes amidetritium label to hemoglobin peptides 14 amino acids or greater in size,without the ability to further sublocalize the label.

Moreover, in Englander's work, there is no appreciation that a suitablyadapted tritium exchange technique might be used to identify the peptideamides which reside in the contacting surface of a protein receptor andits binding partner: his disclosures are concerned exclusively with themapping of allosteric changes in hemoglobin. Furthermore, based on hisoptimization studies (6-11,13), Englander teaches and warns that a pH of2.7 must be employed in both the proteolysis and HPLC steps,necessitating the use of proteases which are functional at these pH's(acid proteases). Unfortunately, acid proteases are relativelynonspecific in their sites of proteolytic cleavage, leading to theproduction of a very large number of different peptide fragments andhence to considerable HPLC separation difficulties. The constraint ofperforming the HPLC separation step at pH 2.7 greatly limits the abilityto optimize the chromatographic separation of multiple overlappingpeptides by varying the pH at which HPLC is performed. Englander triedto work around these problems, for the localization of hemoglobinpeptides experiencing allosteric changes, by taking advantage of thefact that some peptide bonds are somewhat more sensitive to pepsin thanothers. He therefore limits the duration of exposure of the protein topepsin to reduce the number of fragments. Even then the fragments were“difficult to separate cleanly”. They were also, of course, longer (onaverage), and therefore the resolution was lower. He also tried tosimplify the patterns by first separating the alpha and beta chains ofhemoglobin. However, there was a tradeoff: increased tritium loss duringthe alpha-beta separation and the removal of the solvent, preparatory toproteolysis. Englander concludes,

 “At present the total analysis of the HX (hydrogen exchange) behaviorof a given protein by these methods is an immense task. In a largesense, the best strategies for undertaking such a task remain to beformulated. Also, these efforts would benefit from further technicalimprovements, for example in HPLC separation capability and perhapsespecially in the development of additional acid proteases withproperties adapted to the needs of these experiments” (6).

Over the succeeding years since this observation was made, no advanceshave been disclosed which address these critical limitations of themedium resolution tritium exchange technique. It has been perceived thatimprovements to the HPLC separation step were problematic due to theconstraint of working at pH 2.7. The current limited success with smallproteins has made it pointless to attempt similar studies of largerproteins where the problems of inadequate HPLC peptide separation at pH2.7, and imprecision in the ability to sublocalize labeled amides wouldbe greatly compounded. Furthermore, most acid-reactive proteases are ingeneral no more specific in their cleavage patterns than pepsin andefforts to improve the technology by employing other acid reactiveproteases other than pepsin have not significantly improved thetechnique. Given these limitations of medium resolution tritium exchangeart, no studies have been disclosed which utilize proteins with subunitsize greater than 16 kilodaltons.

Allewell and co-workers have disclosed studies utilizing the Englandertechniques to localize induced allosteric changes in the enzymeEscherichia coli aspartate transcarbamylase (22,23). Burz, et al. (22)is a brief disclosure in which the isolated R2 subunit of this enzyme ison-exchanged in tritiated buffer of specific activity 100 mCi/ml,allosteric change induced by the addition of ATP, and then theconformationally altered subunit off-exchanged. The enzyme R2 subunitwas then proteolytically cleaved with pepsin and analyzed for the amountof label present in certain fragments. Analysis employed techniqueswhich rigidly adhered to the recommendations of Englander, utilizing asingle RP HPLC separation in a pH 2.8 buffer.

The authors note difficulty in separating the large number of peptidesgenerated, even from this small protein subfragment, given theconstraints of the Englander methodology. They comment that “theprincipal limitation of this method at present is the separation withcolumns now available”. ATP binding to the enzyme was shown to alter therate of exchange of hydrogens within several relatively large peptidicfragments of the R2 subunit. In a subsequent more complete disclosure(23), the Allewell group discloses studies of the allosteric changesinduced in the R2 subunit by both ATP and CTP. They disclose on-exchangeof the R2 subunit in tritiated water-containing buffer of specificactivity 22-45 mCi/ml, addition of ATP or CTP followed by off exchangeof the tritium in normal water-containing buffer. The analysis compriseddigestion of the complex with pepsin, and separation of the peptidefragments by reverse phase HPLC in a pH 2.8 or pH 2.7 buffer, all ofwhich rigidly adheres to the teachings of Englander. Peptides wereidentified by amino acid composition or by N-terminal analysis, and theradioactivity of each fragment was determined by scintillation counting.In both of these studies the localization of tritium label was limitedto peptides which averaged 10-15 amino acids in size, without higherresolution being attempted.

Beasty, et al. (24) have disclosed studies employing tritium exchangetechniques to study folding of the a subunit of E. Coli tryptophansynthetase. The authors employed tritiated water of specific activity 20mCi/ml, and fragmented the tritium labeled enzyme protein with trypsinat a pH 5.5, conditions under which the protein and the large fragmentsgenerated retained sufficient folded structure as to protect amidehydrogens from off exchange during proteolysis and HPLC analysis. Underthese conditions, the authors were able to produce only 3 proteinfragments, the smallest being 70 amino acids in size. The authors madeno further attempt to sublocalize the label by further digestion and/orHPLC analysis. Indeed, under the experimental conditions they employed(they performed all steps at 12° C. instead of 4° C., and performedproteolysis at pH 5.5 instead of pH in the range of 2-3), it would havebeen impossible to further sublocalize the labeled amides by tritiumexchange, as label would have been immediately lost (off-exchanged) bythe unfolding of subsequently generated proteolytic fragments at pH 5.5if they were less than 10-30 amino acids in size.

Fromageot, et al., U.S. Pat. No. 3,828,102 (25) discloses using hydrogenexchange to tritium label a protein and its binding partner, and Benson,U.S. Pat. No. 3,560,158 and 3,623,840 (26) disclose using hydrogenexchange to tritiate compounds for analytical purposes.

However, none of the methods described in the art are capable oflocalizing the positions of the tritium labels of the labeled proteinsat high resolution, the best resolution in the art generally being onthe order of ≧14 amino acid residues.

Deuterium Exchange Techniques

Fesik, et al (27) discloses measuring by NMR the hydrogen (deuterium)exchange of a peptide before and after it is bound to a protein. Fromthis data, the interactions of various hydrogens in the peptide with thebinding site of the protein are analyzed.

Patterson, et al. (28) and Mayne, et al. (29) disclose NMR mapping of anantibody binding site on a protein (cytochrome-C) using deuteriumexchange. This relatively small protein, with a solved NMR structure, isfirst complexed to anti-cytochrome-C monoclonal antibody, and thepreformed complex then incubated in deuterated water-containing buffersand NMR spectra obtained at several time intervals. The NMR spectra ofthe antigen-antibody complex is examined for the presence of peptideamides which experience slowed hydrogen exchange with solvent deuteriumas compared to their rate of exchange in uncomplexed nativecytochrome-C. Benjamin, et al. (30) employ an identical NMR-deuteriumtechnique to study the interaction of hen egg lysozyme (HEL) withHEL-specific monoclonal antibodies. While both this NMR-deuteriumtechnique, and medium resolution tritium exchange rely on the phenomenonof proton exchange at peptide amides, they utilize radically differentmethodologies to measure and localize the exchanging amides.Furthermore, study of proteins by the NMR technique is not possibleunless the protein is small (less than 30 kD), large amounts of theprotein are available for the study, and computationally intensiveresonance assignment work is completed.

Recently, others (45-50) have disclosed techniques in whichexchange-deuterated proteins are incubated with binding partner,off-exchanged, the complex fragmented with pepsin, and deuterium-bearingpeptides identified by single stage fast atom bombardment (Fab) orelectrospray mass spectroscopy (MS). In these studies, no attempt hasbeen made to sublocalize peptide-bound deuterium within theproleolytically or otherwise generated peptide fragments.

Thus, as is evidenced by the above discussion, there remains a need inthe art for simple and efficient methods whereby the positions oflabeled solvent-accessible peptide amide hydrogens can be localized athigh resolution within the primary amino acid sequence of a polypeptideor protein, as well as simple and efficient methods for studying ormapping the binding sites and/or interaction surfaces of a polypeptideor protein. Accordingly, these are objects of the present invention.

SUMMARY OF THE INVENTION

These and other shortcomings in the art are overcome by the presentinvention, which in one aspect provides methods for the functionallabeling and identification of specific amino acid residues thatparticipate in binding protein-binding partner interactions. The methodsof the invention are particularly suitable for the study of the bindingprotein-binding partner subregions of large (>30 KD) proteins, even insmall quantities.

In one embodiment, the label is tritium and the amount of label on afragment or subfragment is determined by measuring its radioactivity. Ina second embodiment, the label is deuterium and the amount of label on afragment or subfragment is determined by mass spectrometry. The term“heavy hydrogen” is used herein to refer generically to either tritiumor deuterium. In addition, references to tritium apply mutatis mutandisto deuterium except when clearly excluded.

In essence, the binding protein is first tritiated or deuterated underconditions wherein native hydrogens are replaced by the tritium ordeuterium label (this is the “on-exchange” step). Then the bindingpartner is allowed to interact with labeled protein. The binding partneroccludes the binding site and protects the tritium or deuterium labelsof that site from a subsequent “off-exchange”. Thus, after the“off-exchange”, only the binding site residues are labeled. Since thebinding site is normally only a small portion of the molecules, a highersignal-to-background ratio is obtained with this approach than withEnglander's more conventional procedure.

In order to actually identify the labeled residues, one must firstdissociate the complex under slow hydrogen isotope exchange (H³/H¹ orH²/H¹) conditions, since otherwise the labels would leave the bindingsite as soon as the ligand was removed. The binding protein is thenoptionally fragmented (e.g., with an endoprotease such as pepsin), stillunder slow hydrogen exchange conditions, to obtain fragments. Thosefragments which bear label presumably include binding site residues. Atthis point, the resolution of the binding site is no better than thefragment size.

A finer localization of the labels is achieved by analysis ofsubfragments generated by controlled, stepwise, degradation of thebinding protein or of each isolated, labeled peptide fragment (if thebinding protein was optionally fragmented) under slowed exchangeconditions. For the purpose of the present invention, the protein or apeptide fragment is said to be “progressively”, “stepwise” or“sequentially” degraded if a series of fragments are obtained which aresimilar to those which would be achieved by an ideal exopeptidase. Foran ideal exopeptidase, only an end amino acid is removed. Thus, if the namino acids of a peptide were labeled A₁ to A_(n) (the numberingstarting at whichever end the degradation begins), the series ofsubfragments produced by an ideal exopeptidase would be A₂ . . . A_(n),A₃ . . . A_(n), . . . , A_(n−1)−A_(n), and finally An. However, it is tobe understood that while preferably each subfragment of the series ofsubfragments obtained is shorter than the preceding subfragment in theseries by a single terminal amino acid residue, exopeptidases are notnecessarily ideal. Thus, for purposes of the present invention, afragment is said to be “progressively,” “stepwise” or “sequentially”degraded if a series of subfragments is generated wherein eachsubfragment in the series is composed of about 1-5 fewer terminal aminoacid residues than the preceding subfragment in the series. The signalsproduced by the successive subfragments are correlated in order todetermine which amino acids of the fragment in question were labeled.

This procedure was not used in any of the cited references to furtherlocalize the labeling sites, though improved resolution was certainly agoal of the art. The closest the art comes is Englander's generalsuggestions of further fragmentations with another “acid protease”.

The progressive degradation is preferably achieved by an enzyme, andmore preferably by a carboxypeptidase. The need to employ an acidic pHat the time of degradation to minimize tritium losses discourages use ofcarboxypeptidases which are substantially inactivated by the requiredacidic buffers. However, carboxypeptidase-P, carboxypeptidase Y, andseveral other acid-reactive (i.e., enzymatically active under acidconditions) carboxypeptidases are suitable for proteolysis of peptidesunder acidic conditions, even at pH 2.7.

Progressive subfragmentation of purified tritium label-bearing peptidesis performed with acid-reactive carboxypeptidases under conditions thatproduce a complete set of amide-labeled daughter peptides each shorterthan the preceding one by 1-5 carboxy terminal amino acids, andpreferably by a single carboxy-terminal amino acid. HPLC analysis of theseveral members of this set of progressively truncated peptides allowsthe reliable assignment of label to a particular amide position withinthe parent peptide.

Alternatively, the present invention contemplates C-terminal chemicaldegradation techniques that can be performed under “slow hydrogenexchange conditions” e.g., by pentafluoropropionic acid anhydride. Thesensitivity of the technique may be improved by the use of referencepeptide subfragments as HPLC mobility markers.

In general, the art has given insufficient consideration to the problemsof denaturing the binding protein sufficiently to facilitate proteolysisunder slow hydrogen exchange conditions. Pepsin, for example, is muchless active at 0° C. than at room temperature. While pepsin is able toextensively digest hemoglobin that has been denatured by acidic pH at 0°C., certain other binding proteins, such as hen egg lysozyme, are muchmore resistant to denaturation by slow H-exchange conditions, and henceto subsequent pepsin digestion. As a result, many fewer and longerfragments are generated. This complicates the analysis.

In a preferred embodiment, the labeled binding protein is exposed,before fragmentation, to denaturing conditions compatible with slowhydrogen exchange and sufficiently strong to denature the protein enoughto render it adequately susceptible to the intended proteolytictreatment. If these denaturing conditions would also denature theprotease, then, prior to proteolysis, the denatured protein is switchedto less denatured conditions (still compatible with slow H-exchange)sufficiently denaturing to maintain the protein in aprotease-susceptible state but substantially less harmful to theprotease in question.

Preferably, the initial denaturant is guanidine thiocyanate, and theless denaturing condition is obtained by dilution with guanidine HCl.

Disulfide bonds, if present in the binding protein to be digested, canalso interfere with analysis. Disulfide bonds can hold the protein in afolded state where only a relatively small number of peptide bonds areexposed to proteolytic attack. Even if some peptide bonds are cleaved,failing to disrupt the disulfide bonds would reduce resolution of thepeptide fragments still joined to each other by the disulfide bond;instead of being separated, they would remain together. This wouldreduce the resolution by at least a factor of two (possibly more,depending on the relationship of disulfide bond topology to peptidecleavage sites). If the disulfide bonds are not disrupted, furthersublocalization of the tritium-labeled amides within each of thedisulfide-joined peptides would be very difficult, as amino acid removalwould occur, at different times and at different rates, at eachC-terminal of the disulfide linked segments.

The applicant has discovered that water soluble phosphines may be usedto disrupt a protein's disulfide bonds under “slow hydrogen exchange”conditions. This allows much more effective fragmentation of largeproteins which contain disulfide bonds without causing tritium label tobe lost from the protein or its proteolytic fragments (as would be thecase with conventional disulfide reduction techniques which must beperformed at pH's which are very unfavorable for preservation of tritiumlabel).

In another embodiment, peptide amides on the binding protein's surfaceare indirectly labeled by transfer of tritium or deuterium that has beenpreviously attached by hydrogen exchange to the interaction surface ofthe binding partner. This procedure will functionally label receptorprotein amides if they are slowed by complex formation and are also inintimate contact with the binding partner, in the complexed state.Amides that are distant from the interaction surface but slowed inexchange because of complex formation-induced allosteric changes in theprotein will not be labeled.

In another aspect, the present invention provides a method fordetermining which peptide amide hydrogens in a polypeptide or proteinare accessible to solvent. By using the method of the invention, thepositions of solvent-accessible peptide amide hydrogens within apolypeptide or protein can be localized at high resolution, i.e.,typically within 5 or fewer amino acid residues, and in many instancesto within a single amino acid residue.

In the method, solvent accessible peptide amide hydrogens of apolypeptide or protein of interest are on-exchanged by contacting thepolypeptide or protein with heavy hydrogen under conditions wherein thenative solvent-accessible peptide amide hydrogens are replaced withheavy hydrogen (deuterium or tritium), such as, for example,physiological conditions wherein the polypeptide or protein is foldedinto its native conformation. Peptide amide protons that areinaccessible to solvent, such as those that are buried within theinterior of the polypeptide or protein structure or those thatparticipate in intramolecular hydrogen-bonding interactions, do notreadily exchange with the heavy hydrogens in the solvent. Thus, thosepeptide amide hydrogens that are solvent-accessible are selectivelylabeled with heavy hydrogen.

The positions of the labeled peptide amide hydrogens within thepolypeptide or protein can be localized at high resolution byprogressively generating a series of subfragments under conditions ofslowed exchange as previously described, determining which subfragmentsare labeled and correlating the sequences of the labeled subfragmentswith the sequence of the polypeptide or protein to determine whichpeptide amide groups in the polypeptide or protein were labeled, andthus accessible to solvent.

In some embodiments, especially those wherein the polypeptide or proteinof interest is relatively large, the polypeptide or protein mayoptionally be first fragmented (e.g., with an endoprotease or mixture ofendoproteases) under conditions of slowed exchange as previouslydescribed, and the positions of the labels localized to high resolutionby progressively degrading each labeled fragment into a series ofsubfragments, determining which subfragments are labeled and correlatingthe sequences of the labeled subfragments to the sequences of thelabeled fragments and, ultimately, to the sequence of the polypeptide orprotein, as previously described.

In a preferred embodiment, the polypeptide or protein is denatured andany disulfide bonds reduced under conditions of slowed exchange prior tofragmentation and/or subfragmentation.

BRIEF DESCRIPTION OF THE DRAWINGS

FIGS. 1a-1 d depict the results of analysis of tritium associated withhemoglobin (Hgb) fragments produced by pepsin digestion oftritium-exchanged hemoglobin±monoclonal, antibody followed by HPLC inPO₄ buffered solvents, pH 2.7. FIG. 1a: Absorbance (214 nm) tracing ofunlabeled proteolyzed Hgb. FIG. 1b: Hgb on-exchanged for 4 hours,shifted to pH 2.7 and then proteolyzed without off exchange. FIG. 1c:Hgb on-exchanged for 4 hours, mixed with monoclonal antibody β6 and thenoff-exchanged for 40 hours before proteolysis at pH 2.7. FIG. 1d: Hgbon-exchanged for 4 hours and then off-exchanged for 40 hours beforeproteolysis at pH 2.7.

FIG. 2 depicts the results of second dimension separation (HPLC with0.1% Trifluroracetic Acid (TFA) containing solvents) at 0° C. oftritium-bearing rpHPLC fraction from first dimension separation, FIG.1c.

FIGS. 3a-c show the identification of hemoglobin peptides functionallylabeled by interaction with monoclonal antibody β-121. FIG. 3a: Hgbon-exchanged for 4 hours then proteolyzed without a period of offexchange. FIG. 3b: Hgb on-exchanged for 4 hours, mixed with monoclonalantibody β121 and then off-exchanged for 40 hours. FIG. 3c: Hgbon-exchanged for 4 hours and then off-exchanged for 40 hours.

FIGS. 4a-d depict the identification of hemoglobin peptides functionallylabeled by interaction with haptoglobin. FIG. 4a: HPLC optical densitytracing. FIG. 4b: Hgb on-exchanged for 4 hours then proteolyzed withouta period of off exchange. FIG. 4c: Hgb on-exchanged for 4 hours, mixedwith haptoglobin and then off-exchanged for 40 hours. FIG. 4d: Hgbon-exchanged for 4 hours and then off-exchanged for 40 hours.

FIGS. 5a-b show the structure of hemoglobin with peptidic regionshighlighted. FIG. 5A: β6 monoclonal interaction peptides; FIG. 5B: β121monoclonal interaction peptides.

FIGS. 6a-j depict the results of carboxypeptidase-P digestion of β1-14peptide. Tritium-exchange-labeled synthetic β1-14 peptide was digested(0° C.) with carboxypeptidase-P (CP-P) using a range of enzymeconcentrations and digestion times as follows: FIGS. 6a, 6 b—nodigestion; FIGS. 6c, 6 d—2.5 min. digestion with 0.1 mg/mlcarboxypeptidase-P; FIGS. 6e, 6 f—2.5 min. digestion with 0.5 mg/mlcarboxypeptidase-P; FIGS. 6g, 6 h—5 min. digestion with 1.0 mg/mlcarboxypeptidase-P; and FIGS. 6i, 6 j —15 min. digestion with 1.0 mg/mlcarboxypeptidase-P;. HPLC analysis was then performed as in FIGS. 1a-d,but with simultaneous measurement of O.D.214 nm and radioactivity ofcolumn effluent. The positions of the several generated C-terminaltruncated peptide fragments are indicated (numbers 3 through 9).Progressive generation of fragments is observed.

FIGS. 7a-j depict the results of reduction of disulfide bonds at pH 2.7.Tritium-exchange-labeled β1-14 peptide (2 μg at 0° C., pH 2.7) wassupplemented with the peptide endothelin (4 μg), which contains twodisulfide bonds (35), and the mixture incubated without (FIGS. 7a, 7 b)or with (FIGS. 7c-j) 50 mM Tris (2-carboxyethyl) phosphine (TCEP) forvarying times at 0° C. (FIGS. 7a, 7 b (5 mins.); FIGS. 7e, 7 f (5mins.); FIGS. 7g, 7 h (10 mins.); FIGS. 7i, 7 j (30 mins.)), 2 minutesat 22° C. (FIGS. 7c, 7 d). The mixtures were then subjected to HPLC asin FIGS. FIGS. 6a-j. The percent of endothelin that remained unreducedunder each condition is indicated. The fraction of tritium label thatremained attached to the β1-14 peptide is: FIG. 7b (0.35); FIG. 7d(0.25); FIG. 7f (0.38); FIG. 7h (0.35); FIG. 7j (0.32). Fifty percentreduction of endothelium disulfides is accomplished at pH 2.7 with aninsignificant loss of peptide amide-bound tritium from the β1-14peptide. “R” indicates the positions of reduced forms of endothelin.

DETAILED DESCRIPTION OF THE PREFERRED EMBODIMENTS

Biochemical Binding, Generally

Many biological processes are mediated by noncovalent bindinginteractions between a protein and another molecule, its bindingpartner. The identification of the structural features of the twobinding molecules which immediately contribute to those interactionswould be useful in designing drugs which alter these processes.

The molecules which preferentially bind each other may be referred to asmembers of a “specific binding pair”. Such pairs include an antibody andits antigen, a lectin and a carbohydrate which it binds, an enzyme andits substrate, and a hormone and its cellular receptor. In some texts,the terms “receptor” and “ligand” are used to identify a pair of bindingmolecules. Usually, the term “receptor” is assigned to a member of aspecific binding pair which is of a class of molecules known for itsbinding activity, e.g., antibodies. The term “receptor” is alsopreferentially conferred on the member of the pair which is larger insize, e.g., on avidin in the case of the avidin-biotin pair. However,the identification of receptor and ligand is ultimately arbitrary, andthe term “ligand” may be used to refer to a molecule which others wouldcall a “receptor”. The term “anti-ligand” is sometimes used in place of“receptor”.

While binding interactions may occur between any pair of molecules,e.g., two strands of DNA, the present specification is primarilyconcerned with interactions in which at least one of the molecules is aprotein. Hence, it is convenient to speak of a “binding protein” and its“binding partner”. The term “protein” is used herein in a broad sensewhich includes, mutatis mutandis, polypeptides and oligopeptides, andderivatives thereof, such as glycoproteins, lipoproteins, andphosphoproteins, and metalbproteins. The essential requirement is thatthe “binding protein” feature one or more peptide (—NHCO—) bonds, as theamide hydrogen of the peptide bond (as well as in the side chains ofcertain amino acids) has certain properties which lends itself toanalysis by proton exchange.

The binding protein may be identical to a naturally occurring protein,or it may be a binding fragment or other mutant of such a protein. Thefragment or mutant may have the same or different binding characteristicrelative to the parental protein.

Integral membrane proteins are of particular interest, as they aredifficult to crystallize for study by X-ray diffraction. Proteins toolarge to study by NMR methods, e.g., those larger than about 50 kDa, arealso of special interest, particularly if they cannot be characterizedas a composite of two or more separately analyzable domains. Examples ofsuitable proteins include integrins (which are large integral membraneproteins), cell surface receptors for growth factors (including cytokinereceptors), “seven-spanners”, selectin, and cell surface receptors ofthe immunoglobin superfamily (e.g., ICAM-1).

The method of the present invention is especially useful for studyingproteins with discontinuous epitopes, such as certain antibodies,including certain clinically important autoimmune antibodies.

A “binding site” is a point of contact between a binding surface(“paratope”) of the binding protein and a complementary surface(“epitope”) of the binding partner. (When the binding partner is aprotein, the designation of “paratope” and “epitope” is essentiallyarbitrary. However, in the case of antibody-antigen interactions, it isconventional to refer to the antigen binding site of the antibody as the“paratope” and the target site on the antigen as the “epitope”.) Aspecific binding pair may have more than one binding site, and the term“pair” is used loosely, as the binding protein may bind two or morebinding partners (as in the case of a divalent antibody). Moreover,other molecules, e.g., allosteric effectors, may alter the conformationof a member of the “pair” and thereby modulate the binding. The term“pair” is intended to encompass these more complex interactions.

Slowed Hydrogen Exchange Conditions

The present invention contemplates labeling the binding site of abinding protein (or binding partner) with a heavy hydrogen isotope, anddetermining the location of the labels under slowed hydrogen exchangeconditions. “Slowed hydrogen exchange conditions” are hereby defined asconditions wherein the rate of exchange of normal hydrogen for heavyhydrogen at amide hydrogens freely exposed to solvent is reducedsubstantially, i.e., enough to allow sufficient time to determine, bythe methods described herein, the precise amide hydrogen positions whichhad been labeled with heavy hydrogen. The H-exchange rate is a functionof temperature, pH and solvent. The rate is decreased three fold foreach 10° C. drop in temperature. Hence use of temperatures close to 0°C. is preferred. In water, the minimum H-exchange rate is at a pH of2-3. As conditions diverge from the optimum pH, the H exchange rateincreases, typically by 10-fold per pH unit increase or decrease awayfrom the minimum. Use of high concentrations of a polar, organiccosolvent shifts the pH min to higher pH, potentially as high as pH 6and perhaps, with the right solvent, even higher.

At pH 2.7 and 0° C., the typical half life of a tritium label at anamide position freely exposed to solvent water is about 70 minutes.Preferably, the slowed conditions of the present inventions result in ahalf-life of at least 10 minutes, more preferably at least 60 minutes.

Tritium Exchange Embodiments

In one embodiment, the present invention contemplates the followingprocedure for characterization of a binding site or the high resolutionidentification of solvent-accessible peptide amide groups:

A. The phenomenon of hydrogen (tritium) exchange is used to substitute aradioactive probe (tritium) for each of the amide hydrogens on the aminoacids which make up the surface of the receptor protein, including thesurface of the receptor's ligand binding site. This labelling isaccomplished under essentially physiologic conditions by incubating thereceptor protein in solutions containing tritiated water. (Preferably,the water is of high specific activity.)

If one desires to characterize the binding site of the protein, thefollowing steps (B) and (C) are performed.

B. Protein ligand (binding partner) is then added to the on-exchanged(tritiated) receptor protein and allowed to bind to its specific site onthe receptor. Once the ligand has bound to the receptor, hydrogens onthe amino acids which make up the surface of the receptor's binding siteare no longer capable of efficiently interacting with the surroundingaqueous buffer, and further hydrogen exchange is markedly inhibited.

C. The tritiated receptor-ligand complex is then transferred tophysiologic buffers free of tritium. Tritium label on thereceptor-ligand complex is allowed to exchange off the receptor.However, binding complex-dependent hydrogen-bonding between the proteinand binding partner and limited solvent accessibility to theprotein-binding partner interface in the complex are selectiveimpediments to the off-exchange of peptide amide tritium labelsandwiched between the protein and binding partner. After the removal(off-exchange) of tritium from other regions of the protein-bindingpartner complex is substantially finished, the result is thepreferential retention of tritium label at the amides for which hydrogenexchange is slowed by virtue of protein-binding partner interactions,typically amides proximate to amino acids which make up the surface ofthe receptor's ligand binding site. Optionally, the complex may besubjected to limited proteolytic digestion, denaturation and/ordisulfide reduction while off exchange is proceeding, as long as theintegrity of the binding protein: binding partner interaction is notsubstantially perturbed by such maneuvers.

Alternatively, the receptor ligand complex can be tritiated such thatthe peptide amide groups which compose the binding site and/or bindingsurface are not labeled and all other solvent-accessible peptide amidegroups are selectively labeled. Thus, in this alternative embodiment,peptide amide groups which comprise the binding site and/or bindingsurfaces are “functionally labeled” by the absence of tritium.

D. The specific peptide bond amides which bear tritium are thenidentified. This is done by:

(1) shifting the labeled receptor-ligand complex to conditions (e.g.,0-4° C., pH 2.7) which dissociate the complex and at the same time slowdown amide hydrogen exchange.

(2) optionally subjecting the receptor to proteolysis followed byreverse phase (RP) high pressure liquid chromatographic (HPLC)separation (preferably 2-dimensional) of the resulting receptorfragments under continued slow proton exchange conditions. Receptorfragments bearing tritium label are identified, isolated, andcharacterized as to their amino acid sequence, and therefore theirlocation within the primary amino acid sequence of the intact receptor.

Preparation of the binding protein for proteolytic analysis may involve:

(a) trimming off of portions of the protein not required for complexformation;

(b) disruption of disulfide bonds which could complicate the analysis ofthe fragments (see section 5A); and/or

(c) denaturation of the protein to render it more susceptible toproteolytic attack (see section 5B).

Step (a) may be performed before or after switching to slow hydrogenexchange conditions, since it does not cause dissociation of thecontacting surfaces. Steps (b) and (c) are more likely to cause suchdissociation and therefore will more often need to be performed underslow exchange conditions.

(3) determining the location of tritium label within the binding proteinor each labeled peptide fragment from step (2) by subfragmenting thelabeled binding protein or peptides (e.g., with acid-reactivecarboxypeptidases or tritium-exchange-compatible chemical methods) underslow proton exchange conditions and characterizing the labelledsubfragments. For example, the identity of each of the subfragments maybe determined by amino acid analysis, peptide sequencing, or bycomparison of their mobility with synthetic HPLC mobility markerpeptides, and the amount of tritium label attached to each subfragmentdetermined by scintillation counting. As each carboxy-terminal aminoacid of the functionally labeled binding protein or peptide fragment issequentially cleaved off by the carboxypeptidase, the nitrogen whichformed the slowly-exchanging peptide amide in the intact peptide bond isconverted to a rapidly exchanging secondary amine, and any tritium labelat that nitrogen is lost from the peptide within seconds, whereas allother amide bond tritium remains in place. A stepdown in radioactivityfrom one subfragment to the next smaller one indicates that the amidejust altered had been labeled with tritium.

In this manner, the precise location, within the protein, of eachpeptide amide that is functionally labeled by virtue of itssolvent-accessibility and/or its interaction with its binding partner isdetermined. Inferentially, in this manner, the precise amino acids whichmake up the surface of the receptor and/or the surface of the receptor'sbinding site are then known. Studies may be performed to quantify theexchange rates of each of the labeled amides identified above bothbefore and after complex formation with binding partner. This allowscalculation of the magnitude of exchange slowing experienced by each ofthese amides consequent to complex formation, and allows optimization ofon and off exchange times.

E. Parallel studies may be performed in which the cognate bindingpartner is on-exchanged with tritium, complexed with receptor protein,off-exchanged as a binding partner-protein complex and slowed amides inthe binding partner identified as above. This procedure results in theidentification of the subregions of the binding partner which interactwith the protein.

F. The knowledge of the identity of the precise contact peptides in bothreceptor and ligand may be combined with additional structuralinformation provided by the invention (identification of peptide amidesof the protein and binding partner which are likely to directly formhydrogen bonds between protein and binding partner upon complexformation) to produce models for the complementary 3-dimensionalstructures of the receptor and ligand interaction surfaces. These modelsmay then be used as the basis of the design and production ofappropriate peptide and peptidomimetic drugs.

The individual steps of this procedure will now be considered in greaterdetail.

1. On-Exchange

The protein under study is incubated in buffer supplemented withtritiated water (³H₂O), preferably of high specific activity. Thisresults in the time dependent reversible incorporation of tritium labelinto every peptide amide on the surface of the protein, including its(potential) ligand binding subregion, through the mechanism of protonexchange.

Any physiologic buffer appropriate for the interaction of the proteinwith its binding partner may be utilized (with no constraints imposed onbuffer pH or temperature). Suitable buffers include phosphate bufferedsaline (PBS), 0.15 mM NaCl, 10 mM PO₄, pH 7.4 PBS. The use of smallincubation volumes (0.1-10 μl) containing high concentrations ofreceptor protein (10-100 mg/ml) is preferred.

The necessary level of tritiation (and hence the concentration oftritium in the buffer) is dependent on the total amount of proteinavailable for analysis. For analysis of 1 mg protein, at least 10 Ci/mlis desirable; for 0.1 mg, 100 Ci/ml, and for 0.01 mg, 1000 Ci/ml. (Puretritiated H₂O is about 2500 Ci/ml.) For most applications, the tritiatedwater will be 50-500 Ci/ml. Without the use of these high specificactivities, studies of proteins which are available in limited quantitywould be much more difficult. (Even higher specific activity (e.g.,500-1,500 Ci/ml) may be used in the invention, but radiation safetyconsiderations necessitate performance of such on- and off-exchangeprocedures in specialized facilities, such as are available in thetritium laboratory provided by the National Tritium Facility, LawrenceBerkeley Laboratories, University of California, Berkeley.)

It should be noted that with customary levels of tritium, only a smallpercentage of the binding protein molecules will be tritiated at anygiven exposed position. All that is required is that substantially eachof the exposed amide hydrogen atoms be replaced in a detectable (byradiation counting) number of the binding protein molecules.

It is not necessary that the tritium exchange analysis rely on only asingle choice of “on-exchange” time. Rather, the skilled worker maycarry out the experiment using a range of on-exchange times, preferablyspanning several orders of magnitude (seconds to days) to allowselection of on-exchange times which allow efficient labeling of thevarious peptide amides present in the protein, which will become slowedin their exchange rate consequent to the interaction of the protein toits binding partner, and at the same time minimize background labelingof other amide positions after off-exchange is completed (see section 10below).

2. Receptor-Binding Partner Complex Formation

After a suitable period of tritium on-exchange, the protein's bindingpartner is added to the tritiated protein-buffer solution and the twoallowed to form a binding complex. The binding partner is preferablyadded in quantities sufficient to produce saturation binding to theprotein (usually equimolar amounts) and at high concentrations (e.g.,10-100 mg/ml) to maximize the rate and extent of binding. To minimizetritium labeling of the added binding partner by proton exchange(important when utilizing short on-exchange times), ³H₂ 0 in the bufferis preferably diluted with tritium-free buffer (10-1000 fold dilution)within 0-100 seconds of binding partner addition. Additionalmanipulations detailed below may be used at this step to furtherminimize incorporation of tritium label into the binding partner.

3. Off-Exchange

The tritiated protein-binding partner complex is then transferred tophysiologic buffers identical to those employed during on-exchange, butwhich are substantially free of tritium. Tritium label on the proteinthen exchanges off the protein at rates identical to its on-exchangerate everywhere except at amides which have been slowed in theirexchange rate by virtue of the interaction of protein with bindingpartner. With sufficient off-exchange time, the result is the specificretention of tritium label at each of the peptide amide bonds whichoccur between the amino acids which make up the surface of the protein'sbinding site for the binding partner. We refer to this process as acomplex formation-dependent functional labeling of the protein withtritium. At least 90%, more preferably, at least 95%, 96%, 97%, 98% or99% or more, of on-exchanged tritium label at other sites isoff-exchanged from the protein.

In general, off-exchange is allowed to proceed for 5 to 50 times, morepreferably about 10 times longer than the on-exchange period, as thisallows off-exchange from the protein of greater than 99% of theon-exchanged tritium label which has not experienced a slowing ofexchange rate subsequent to the protein's interaction with bindingpartner. Preliminary studies may be performed with the protein andbinding partner to determine the on and off exchange times whichoptimize the signal (tritium remaining in functionally labeled amides)to noise (tritium remaining in background amides) ratio (see section 8).

In preferred embodiments, the off-exchange procedure may be performedwith the use of Sephadex G-25 spin columns prepared and utilized asdescribed in Example 1 (below), by G-25 column chromatography asdescribed by Englander (6,19) or by use of perfusive HPLC supports thatallow rapid separation of peptide/protein from solvent (Poros® columns,PerSeptive Biosystems, Boston, Mass.). Use of the G-25 spin columnsallows the separation of the complex from greater than 99.9% of buffertritium. Residual buffer tritium and tritium off-exchanged from thecomplex may optionally be further removed by dialysis of the complexagainst tritium free buffer during off exchange.

Alternatively, complex formation and off-exchange can be accomplished byfirst reacting the on-exchanged protein-buffer mixture with bindingpartner which has been covalently attached to a solid support (e.g.binding-partner-Sepharose), allowing the on-exchanged protein to complexto the solid-phase binding partner, followed by washing of thesepharose-binding partner-protein conjugate with tritium free buffer.Alternatively, soluble protein-binding partner complexes may be formedas above, and captured with a solid phase adsorbent that can bind toeither the protein or binding partner component of the complex (e.g.Sepharose with covalently attached antibodies specific for protein orbinding partner).

Most protein-ligand binding interactions that will be probed with thistechnique are reversible reactions: binding partner will dissociate fromand rebind to the protein during the off-exchange period, and during thebrief intervals where the protein's binding site is unoccupied withbinding partner, proton off-exchange proceeds at the unprotected rate.It is therefore important to minimize the time that the binding site isunoccupied. In a preferred embodiment, this is accomplished by havingboth receptor and binding partner present at high concentration, e.g.,at least mg/ml concentrations, up to 100 mg/ml concentrations eachthroughout the off-exchange period, and performing the on and offexchange reactions at temperatures at or below room temperature,preferably 4° C.

4. Trimming of the Binding Protein (optional)

Prior to dissociation of the complex, e.g., during the off-exchangeperiod, which typically lasts hours to days, the complex may optionallybe chemically or enzymatically treated to produce the smallest fragmentof protein which is still capable of remaining tightly bound to thebinding partner, and this residual “trimmed” complex isolated. Removalof portions of the protein not essential for continued complex formationwill decrease the number of background peptides generated during thesubsequent acid proteolysis of the trimmed complex (Section 6). Thispre-digestion and purification can be performed with a wide variety ofproteases (e.g. trypsin, pronase, V-8 protease chymotrypsinproteinase-K) as well as certain chemical agents (e.g., cyanogenbromide, iodosobenzoic acid), and under virtually any conditions ofinduced partial protein denaturation (e.g. urea, guanidinium chloridesodium dodecyl sulfate, non-ionic detergents, reductants such as2-mercaptoethanol, dithiothreitol), ionic strength, temperature, timeand pH which do not substantially dissociate the contacting surfaces ofthe protein-binding partner complex. Excessive digestion efforts whichresult in dissociation of these surfaces from each other will cause alarge fraction of functional tritium label to be immediatelyoff-exchanged, as greater than 50% of peptide amides in the dissociatingsurfaces will have exchange half-lives of less than 1 minute atapproximately 15 pH 7. The goal is to generate and isolate a fragment ofthe protein, preferably 15-100 kD in size more preferably 15 kD, whichremains attached to the binding partner. Often “ligand stabilization” ofproteins which are proteolysed while bound to binding partner allows thecontinued binding of the protein fragments to partner.

Preliminary studies may be performed with the off-exchanged complex todetermine conditions which result in a suitably trimmed protein-bindingpartner complex. In a preferred embodiment, the quantity of residualtritium functionally bound to the intact off-exchanged complex is firstdetermined by measurement of tritium which migrates with the void volume(Mr>10,000 kD) on a G-25 spin column (pH 7.4). Aliquots of the complexare then subjected to varied fragmentation conditions, and the fractionof tritium label which remains attached to polypeptides under eachdigestion condition (migrates with G-25 void volume) determined. Theproteolytic products of the most vigorous digestions which “release”less than 5% of complex-associated tritium are (as per Section 5)adjusted to pH 2.7, 0° C., subjected to RP-HPLC at pH 2.7, 0° C., andpeptides/protein fragments which bear label identified, isolated, andtheir molecular weights determined by SDS-PAGE. The labeled proteolyticproducts produced in these limited digests are likely to be largepolypeptides, and therefore RP-HPLC supports suitable to thepurification of such peptides (C-4, phenyl columns) are utilized.Alternatively, when solid-phase adsorbents are used for complexformation/off-exchange (step 3), proteolysis as above, now of the solidphase binding partner-protein complex, is allowed to proceed asextensively as possible without release from the solid support ofgreater than 5% functionally attached tritium. The predigestedprotein/complex is then released from the immunoadsorbent withdenaturants including a shift to pH 2.7, and the predigested proteinfurther proteolysed with pepsin other acid reactive proteases.

A binding protein may also be trimmed earlier, e.g., before“on-exchange” or before complex formation, provided that the trimmedprotein binds the partner sufficiently similarly to the original proteinto be of interest.

5. Switch to Slow Amide Hydrogen Exchange Conditions

The protein-binding partner complex (or predigested complex—see Step 4)or selectively labeled protein in the case of alternative embodiments ofthe invention wherein the positions of solvent accessible peptide amideprotons are to be determined in the absence of a binding partner, isthen shifted to conditions of temperature and pH which greatly slow thehalf life of peptide amide hydrogen exchange, and essentially “freeze”in place the tritium labels. In a preferred embodiment, the complex isshifted to 0° C., and pH 2.7 conditions under which the half life ofexchange of peptide amide label in fully denatured peptides is at least70 minutes. The label will be sufficiently held in place under theseconditions so that several rounds of proteolytic fragmentation, HPLCseparation, and tritium quantification can be performed withoutunacceptable loss of label.

For some binding proteins, switching to slow hydrogen exchangeconditions is sufficient to cause dissociation of the complex. If not, adissociating agent, such as a chaotropic agent may be added.

5A. Disruption of Disulfide Bonds (optional)

High resolution localization of tritium label-bearing amides requiresthe proteolytic generation of peptides less than approximately 15-20amino acids in size under conditions which allow the label to remain inplace (e.g., 0° C., pH 2.7). The ability of any protease to fragment aprotein or peptide is limited by the accessibility of the protease tosusceptible peptide bonds. While denaturants such as acidic pH, urea,detergents, and organic co-solvents can partially denature proteins andexpose many otherwise structurally shielded peptide bonds, pre-existingdisulfide bonds within a protein can prevent sufficient denaturationwith these agents alone. In conventional protein structural studies,disulfides are usually cleaved by reduction with 2-mercaptoethanol,dithiothreitol, and other reductants which unfortunately require a pHgreater than 6 and elevated temperature for sufficient activity, and aretherefore not useful for the reduction of disulfides at pH 2.7 or below.For this reason, the tritium exchange art has not attempted any form ofdisulfide bond disruption, has for the most part been restricted to thestudy of proteins without intrinsic disulfide bonds, and has acceptedthe low resolution achievable without disulfide bond disruption. Theapplicant has recognized and demonstrated that acid-reactive phosphinessuch as Tris (2-carboxyethyl) phosphine (TCEP) (31-36) can be used todisrupt disulfides under the acidic pH and low temperature constraintsrequired for tritium exchange analysis (see FIGS. 7a-j). Thesemanipulations disrupt these associations and at the same time continueto produce a markedly slowed proton exchange rate for peptide amideprotons.

5B. Protein Denaturation. (optional)

In previous studies by Englander et al. and others, employing mediumresolution tritium exchange, proteolytic fragmentation oftritium-labelled proteins under slowed-exchange conditions wasaccomplished by shifting the protein's pH to 2.7, adding highconcentrations of liquid phase pepsin, followed by brief (10 min.)incubation at 0° C. With the proteins studied by Englander et al. simplyshifting pH from that of physiologic (7.0) to 2.7 was sufficient torender them sufficiently denatured as to be susceptible to pepsinproteolysis at 0° C. Furthermore, these proteins, in general, did notcontain disulfide bonds that interfered with effective denaturation bysuch (acid) pH conditions or contain disulfide bonds within portions ofthe protein under study with the technique. The applicant has found thatother proteins (for example hen egg lysozyme) are negligibly denaturedand are not substantially susceptible to pepsin proteolysis whencontinuously incubated at comparable acidic pH and depressed temperature(10-0° C.). This is the consequence of the existence of a thermalbarrier to denaturation for many proteins incubated in many denaturants;i.e., denaturation of proteins at lower temperatures (10-0° C.) is ofteninefficient and a slow process, incompatible with the requirement ofmedium resolution tritium exchange techniques that manipulations beperformed rapidly, such that the attached tritium label is substantiallyretained at functionally labelled amides of the binding protein.

The applicant has discovered that such proteins become extraordinarilysusceptible to pepsin proteolysis at 0° C. when they are treated withthe sequential denaturation procedure described below. Furthermore, theapplicant has discovered that although TCEP can effect the reduction ofdisulfide bonds in proteins at 0° C. and pHs in the range of 2-3, it isrelatively inefficient at doing so under these conditions and becomesmuch more efficient at effecting reduction at a pH of 5.0 or greater.Conditions can be arranged to greatly increase the efficiency ofTCEP-mediated reduction while at the same time preserving slow exchangeconditions. This is accomplished by simultaneously denaturing theprotein with guanidine thiocyanate, employing very high concentrationsof TCEP and raising the pH of the solution to 5.0. While this pH wouldnormally produces an unacceptable 100-fold increase (as compared to thatat pH 2.7) in the rate of loss of tritium from the labelled protein, theelevated pH-induced increase in the rate of tritium loss issubstantially offset by limiting the water content of the incubationmixture (and thereby markedly slowing the rate of tritium loss) when theprotein is being reduced at pH 5.0, and the solution pH then is shiftedback to pH 2.7 once reduction is complete. The result is effectivereduction of proteins at a pH of 5 and 0° C. with substantially completeretention of tritium label on the binding protein.

The denatured (or denatured and reduced) protein solution is then passedover a pepsin-agarose column, resulting in efficient and rapidfragmentation of the protein (in≦1 min.). The fragments can be, andusually are, immediately analyzed on RP-HPLC without unnecessarycontamination of the peptide mixture with the enzyme pepsin or fragmentsof the enzyme pepsin. Such contamination is problematic with thetechnique as taught by Englander, et al., as high concentrations ofpepsin (often equal in mass to the protein under study) are employed, toforce the proteolysis to occur sufficiently rapidly at 0° C.

While proteins are often subjected to purposeful denaturation withagents other than a pH shift prior to digestion with pepsin, this hasnever been done at depressed temperatures (10-0° C.) before, and theapplicant has discovered that while guanidine thiocyanate at theindicated concentrations is sufficient to suitably denature and rendersusceptible to pepsin proteolysis proteins at 10-0° C., several otherstrong denaturants, including urea, HCl, sodium dodecyl sulfate (SDS)and guanidine HCl, were, at least when used alone, unable to adequatelydenature lysozyme at these low temperatures. However, the concentrationsof guanidine thiocyanate required for such denaturation are incompatiblewith pepsin digestion; i.e., they denature the pepsin enzyme before itcan act on the denatured binding protein. When the guanidine thiocyanateis removed (at 10-0° C.) from the solution after protein denaturationhas been accomplished in an attempt to overcome this inhibition ofpepsin activity, the protein rapidly refolds and/or aggregates, whichrenders it again refractory to the proteolytic action of pepsin. Theapplicant has discovered that if proteins are first denatured in ≧2Mguanidine thiocyanate at 0° C. and the concentration of thiocyanate thenreduced to ≦2M while at the same time the guanidine ion is maintained≧2M (by diluting the guanidine thiocyanate into guanidinehydrochloride), the denatured protein remains in solution, remainsdenatured, and the enzyme pepsin is efficiently proteolytically activeagainst the denatured protein in this solution at 0° C. The stability ofpepsin-agarose to this digestion buffer is such that no detectibledegradation in the performance of the pepsin column employed by theapplicant has occurred after being used to proteolyze more than 500samples over 1 years. No pepsin autodigestion takes place under theseconditions.

Denaturation without concomitant reduction of the binding protein may beaccomplished by contacting it (at 0-5° C.) with a solution containing ≧2molar guanidine thiocyanate pH 2.7, followed by the addition of an equalvolume of 4 molar guanidine hydrochloride pH 2.7.

Denaturation with disulfide reduction may be accomplished by contactingthe binding protein with a solution containing ≧2 molar guanidinethiocyanate, 0.3-0.7 molar TCEP, 5-20% H₂O (by volume), with the balanceof volume being acetonitrile, dimethyl sulfoxide, or other watermiscible nonaqueous solvent in which the denaturant (e.g. guanidinethiocyanate) and disulfide bond disrupting agent (e.g., TCEP), if used,remain soluble at substantially these concentrations, and such that thesolvent system does not freeze at the “slow exchange” temperature. ThepH of the solution is preferably in the range of 4.8-5.2, optimally 5.0.After this incubation, 2 volumes of a 2.5 molar guanidine hydrochloridesolution is added, with the pH and buffering capacity of the solutionsuch as to achieve a pH of 2.7 in the final mixture.

Denatured (with or without reduction) binding protein is then passedover a column composed of insoluble (solid state) pepsin, whereby duringthe course of the passage of such denatured or denatured and reducedbinding protein through the column, it is substantially completelyfragmented by the pepsin to peptides of size range 1-20 amino acids at0° C. and at pH 2.7. The effluent from this column (containingproteolytically-generated fragments of binding protein) is directly andimmediately applied to the chromatographic procedure employed toseparate and isolate protein fragments, preferably analyticalreverse-phase HPLC chromatography.

It should be noted that denaturants, besides rendering the bindingprotein more susceptible to proteolysis, also help dissociate it fromits partner.

6. Generation of Peptide Fragments (Optional)

To ultimately localize the protein's amides which are functionallylabeled with tritium, small peptides bearing the retained tritium label(preferably, 5-25 amino acids in size) are optionally proteolyticallygenerated from labeled protein and separated from the many otherunlabeled peptides generated by fragmentation of the protein, all underconditions which minimize off-exchange of amide tritium from thepeptide. Small peptides have little secondary structure and thereforetheir amides are free to exchange with solvent hydrogen. If tritiumlabel is to remain in place on such peptides, proteolysis andpurification (e.g., RP-HPLC) conditions must be adjusted to slow suchoff-exchange.

The labeled and dissociated binding protein is therefore fragmentedunder slow H-exchange conditions, e.g., by proteolysis with highconcentrations of a protease which is stable and active with theaforementioned conditions (e.g., pH 2.7, 0° C.). Suitable acid tolerantproteases include pepsin (19), cathepsin-D (37) Aspergillus proteases(37a-37c), thermolysin (38) and mixtures of these proteases. In apreferred embodiment, pepsin is used, preferably at a concentration of10 mg/ml pepsin at 0° C. pH 2.7 for 5-30 minutes, preferably 10 minutes.

Other physical and chemical fragmentation methods may be used providedthey are (1) are compatible with slow H-exchange conditions, (2) do notcause shifts in the positions of the amide labels, and (3) produce areasonable number of fragments from the protein of interest.

Preferably, prior to fragmentation of the binding protein, bindingpartner (if susceptible to the fragmenting agent) is removed, so as notto complicate purification with binding partner fragments.

6A. Purification of Fragments

As acid proteases in general have very broad cleavage specificity, theyfragment the protein into a very large number of different peptides. Inmost protein-binding partner systems studied by tritium exchange, it islikely that the interacting binding surfaces will contain roughly 10-20tritium labeled peptide amide which upon proteolysis will result inapproximately 1-5 label-bearing peptides, the precise number dependingon the inherent fragmentation mode of the protein under study with theproteases utilized. The number of “background,” non-labeled peptides(derived from regions of the protein and binding partner that do notparticipate in the binding interaction) generated by the fragmentationprocedure will be a direct function of the size of the protein.Background peptides will be present in the proteolytic digest in numbers10-1,000 times greater than will be functionally labeled peptides whenproteins with sizes in the range of 30-200 kD are proteolyzed.

This large number of background peptides causes two difficulties: First,they must all be cleanly separated from the functionally labeledpeptides to allow identification of the label-bearing peptides. Second,background peptides contain small amounts of tritium label and eventhough the amount of label per background peptide is generally less than1% of that of functionally labeled peptides, background peptides arepresent in much greater amounts and are likely to obscure the presenceof functionally labeled peptides and analytical separation.

Given these considerations, only proteins less than 30 kD in size havebeen successfully characterized in the past by medium resolution tritiumexchange. Upon acid proteolysis of larger proteins, so many differentfragments would be obtained that individual fractions obtained on asingle HPLC separation performed at pH 2.7 would be unacceptablycontaminated with background peptides.

Any method of purifying the fragments which is capable of resolving themixture while maintaining slow H exchange condition is acceptable. Thepreferred method is high pressure liquid chromatography (HPLC),especially in reverse phase (RP). (An alternative method is that of massspectroscopy.)

The art has overstated the sensitivity of the tritium label to pH.Englander (10) reported that at 0° C., the tritium label was most stable(when the tritiated protein was placed in an untritiated aqueous buffer)at pH 2.7, and that the rate of off-exchanged increased rapidly (10 foldper pH unit) as one moved away from that pH. Surprisingly, Applicantfound that at 0° C., the label was sufficiently stable to permitanalysis even at a pH of 2.1. While the acceptable pH range will varywith temperature, and the choice of solvent (the optimal pH increases ifa polar nonaqueous solvent is introduced), the fact remains that pH waspreviously considered to be essentially fixed. Since the tritium labelis stable over a broader pH range, such as 2.1-3.5, it is possible todepart from Englander's recommended pH of 2.7 in seeking HPLC conditionswhich result in effective separation of the peptide fragments.

When the binding molecules are large, so many different fragments areobtained after proleolytic digest that some of the individual peaks on asingle HPLC separation, even at optimized ph, may be heterogeneous.

RP-HPLC resolution of co-migrating multiple peptides may be greatlyimproved by resorting to a two-dimensional RP-HPLC separation in whichtwo sequential RP-HPLC separations are performed at substantiallydifferent pH's, e.g. 2.7 and 2.1.

A two-dimensional HPLC separation allows high efficiency purification oftritium label bearing-peptides from the enormous number of unlabeledpeptides generated by peptic fragmentation of large proteins.Two-dimensional separation of molecules is known in the chromatographicart. However, despite frequent complaints in the Tritium exchangeliterature about resolution problems, 2D separations have not beenemployed previously in connection with Tritium exchange.

In a preferred embodiment of the invention, tritium-labeled proteinfragments are first separated by means capable of sufficiently resolvingthe fragments, such as by RP-HPLC (utilizing any of a number ofpotential chromatographic support including C4, C18, phenol and ionexchange, preferably C18). This separation may be performed at pH2.1-3.5 and at 4-0° C., more preferably, at pH 2.7 and 0° C., which mayaccomplished by employment of any buffer systems which operate at thispH, including citrate, chloride, acetate, more preferably phosphate.Peptides are eluted from the reverse phase column with a similarlybuffered gradient of polar co-solvents including methanol, dioxane,propanol, more preferably acetonitrile. Eluted peptides are detected byon-line ultraviolet light absorption spectroscopy performed atfrequencies between 200 and 300 nm, preferably 214 nm. Tritium label isdetected by scintillation counting of a sampled fraction of the HPLCcolumn affluent. Peptides bearing label that has been specificallyprotected from off-exchange by complex formation with binding partnerare identified by comparing the specific activity of each labeledpeptide to the specific activity of the same peptide prepared fromprotein subjected to identical on/off exchange, proteolysis and HPLCconditions, but which have been off-exchanged without added bindingpartner.

HPLC fractions containing peptides with such functionally labeled amidesare then subjected to a second dimension RP-HPLC separation which may beperformed at pH 2.1-3.5 and 4-0° C., more preferably, at pH 2.1 and 0°C., accompanied by any buffer systems which operates at this pH,including citrate, chloride, acetate, phosphate, more preferably TFA(0.1-0.115%). Peptides are eluted from their reverse phase column with asimilarly buffered gradient of polar co-solvents including methanol,dioxane, propanol, more preferably acetonitrile. Eluted peptides aredetected, tritium measured and functionally labeled peptides identifiedas in the first HPLC dimension described above. Functionally labeledpeptides are isolated (collection of the appropriate fraction of columneffluent), water, acetonitrile, and TFA removed by evaporation, and theremaining purified peptides each characterized as to its primary aminoacid structure by conventional techniques, e.g., amino acid analysis ofcomplete acid hydrolysates or gas-phase Edman degradationmicrosequencing. Reference is then made to the previously known aminoacid sequence of the intact protein to infer the location of thetritium-labeled peptides within the intact protein's primary sequence.Employment of TFA buffer in the second dimension has the additionaladvantage that no residual salt (i.e. phosphate) remains after solventevaporation. Residual phosphate frequently interferes with the chemicalreactions required for amino acid analysis and Edman degradation, aproblem obviated by the use of volatile TFA in the second dimensionbuffer.

Most preferably, proteolytic digests are first separated at pH 2.7 inphosphate buffered solvents and each eluted peptide peak fraction whichcontains tritium-labeled amides is identified, collected, and thensubjected to a second HPLC separation performed in trifluoracidic acid(TFA)-buffered solvents at pH 2.1.

7. High Resolution Sublocalization of Labeled Amides WithinLabel-Bearing Peptides

To routinely localize peptide amide tritium label to the single aminoacid level, applicant systematically cleaves every peptide bond withinthe labeled protein or a purified label-bearing peptide fragment. SlowH-exchange conditions must be used for this proteolysis as the smallpeptides generated have no stable conformational structure and rapidloss of tritium label from the amides would occur if rates of exchangewere not slowed, e.g., by ambient acidic pH.

Most known acid-reactive proteases cleave peptides in a basicallynonspecific manner similar to that of pepsin; studies employing otherpepsin-like proteases have not proved to be of significant utility inincreasing resolution of labeled amides.

A special class of acid-reactive proteases, the carboxypeptidases, areable to generate all required subfragments of labeled protein orpepsin-generated peptides in quantities sufficient for high resolutiontritium localization. Many carboxypeptidases are active at pH 2.7 andsequentially cleave amino acids from the carboxy terminus of peptides.Such enzymes include carboxypeptidase P, Y, W, and C (39). Whilecarboxypeptidases have been used for limited carboxy-terminal sequencingof peptides, often at pH in the range of 2.7 (40), their use in tritiumexchange techniques has not been disclosed. The need to minimize tritiumlosses forbids the use of carboxypeptidases which are inactive in acidic(pH 2.7) buffers, such as carboxypeptidases A and B. However,carboxypeptidase-P, Y, and several other acid-reactive carboxypeptidases(W,C) are suitable for proteolysis of peptides under acidic conditions(39). The tritium exchange art has failed to recognize the utility ofcarboxypeptidases to tritium exchange studies, possibly because thecarboxypeptidases are even more nonspecific in the types of peptidebonds they cleave than are pepsin-like proteases and therefore mighthave been thought to result in inadequate recovery of any singlesubfragment.

Furthermore, chemical procedures employing pentafluoropropionicanhydride can produce sets of C-terminal-truncated peptide fragmentsunder slowed amide exchange conditions (see below, 41,42).

In the preferred embodiment, tritium-exchange-labeled proteins arenonspecifically fragmented with pepsin or pepsin-like proteases, theresulting tritium-labeled peptides isolated by two-dimensional HPLC andthese in turn exhaustively subfragmented by controlled, step-wisedigestion with acid-(i.e., enzymatically active under acidic conditions)exopeptidases and/or by chemical means (see below). These digests arethen analyzed on RP-HPLC performed at 0° C. in TFA-containing buffers(pH 2.1) and each of the generated subfragments (typically 5-20) is thenidentified. The identity of each of the several subfragments maybedetermined by any suitable amino acid analysis, peptide sequencing, orthrough the use of synthetic HPLC mobility marker peptides, and theamount of tritium label attached to each subfragment truncated peptidedetermined by scintillation counting. In this manner, the preciselocation, within the protein, of each peptide amide that is functionallylabeled with tritium by virtue of its interaction with binding partneris determined. By consideration of the tritium content of each of theidentified subfragments the amide hydrogens which had been replaced bytritium during the “on-exchange” step may be inferred. It should benoted that the purpose of the carboxypeptidase treatment is to generatethe subfragments; the method does not require use of carboxypeptidase tosequence the fragments or subfragments. Preferably, the sequence of thebinding protein, or at least of the material portion thereof, is knownprior to commencement of the present method. However, it may bedetermined at any time, even after the subfragmentation, although thedata gleaned from the subfragmentations cannot be properly interpreteduntil the sequences of a least the source is known.

Controlled sequential carboxy-terminal digestion of tritium-labeledpeptides with carboxypeptidases can be performed under conditions whichresult in the production of analytically sufficient quantities of a setof carboxy-terminal truncated daughter peptides each shorter than thepreceding one by from one to about 5 carboxy-terminal amino acidresidues, preferably by a single carboxy-terminal amino acid. As eachcarboxy-terminal amino acid of the functionally labeled peptide issequentially cleaved by the carboxypeptidase, the nitrogen which formedthe slow-exchanging peptide amide in the intact peptide bond isconverted to a rapidly exchanging secondary amine, and any tritium labelat that nitrogen is lost from the peptide within seconds, even at acidicpH. A difference in the molar quantity of tritium label associated withany two sequential subpeptides implies that label is localized at thepeptide bond amide which differs between the two subpeptides.

In a preferred embodiment, synthetic peptides are produced (by standardpeptide synthesis techniques) that are identical in primary amino acidsequence to each of the functionally labeled pepsin-generated peptidesidentified in Step 6. The synthetic peptides may then be used inpreliminary carboxypeptidase digestion (pH 2.7, 0° C.) and HPLC (inTFA-buffered solvents) studies to determine; 1) the optimal conditionsof digestion time and protease concentration which result in theproduction and identification digestion on all possible carboxypeptidaseproducts of the peptide under study; and 2) the HPLC elution position(mobility) of each carboxypeptidase-generated subfragment of syntheticpeptide.

To facilitate this latter procedure, a set of reference peptides may beproduced consisting of all possible carboxyterminal truncated daughterpeptides which an acid carboxypeptidase could produce upon digestion ofa “parent” peptide. These serve as HPLC mobility identity standards andallow the deduction of the identity of daughter peptides actuallygenerated by carboxypeptidase digestion. Certain daughter peptides maybe enzymatically produced in quantities insufficient for direct aminoacid analysis or sequencing, but their HPLC mobility can be measured andcompared to that of the synthetic peptides. Peptides can be detected andquantified by standard in-line spectrophotometers (typically UVabsorbance at 200-214 nM) at levels well below the amounts needed foramino acid analysis or gas-phase Edman sequencing.

After these preliminary studies, the pepsin-generated HPLC isolated,functionally labeled peptide (prepared in Step 6) is thencarboxypeptidase digested and analyzed under the foregoingexperimentally optimized conditions, the identity of each fragmentdetermined (by peptide sequencing or by reference to the mobility ofreference peptide mobility marker) and the amount of tritium associatedwith each peptide subfragment determined.

Alternatively, a chemical technique may be used for the successivecarboxy terminal degradation of peptides under slowed tritium exchangeconditions. Tritium-labeled peptides in HPLC buffers are held at −35° C.and solvents removed by cryosublimation (40a, 40b; vacuum at 1-20millitorr, solvents collected in a liquid nitrogen trap). The driedpeptide is then reacted with vapor phase pentafluoropropionic acidanhydride (PFPA) as described in (54, 55) except that the peptidetemperature is kept at −35° C. for times up to 3 hours. PFPA is thenremoved by vacuum and the fragmented peptide made to 50 mM PO₄ pH 2.7,and analyzed by HPLC.

In general, the known aminopeptidases are not able to sequentiallydegrade a peptide under slow hydrogen exchange conditions. However, ifan acid-reactive aminopeptidase is discovered in nature, or produced bymutation of a known aminopeptidase, there is no reason that anaminopeptidase can not be used in place of the presently preferredcarboxypeptidase. In that event, the stepwise degradation will begin atthe N-terminal, rather than the c-terminal, of each analyzed peptidefragment.

It should be noted that by using polar, nonaqueous high concentrationsof cosolvents to shift the pH_(min) of the H-exchange rate, a greatervariety of reagents may be used than would otherwise be the case. Acosolvent of particular interest in this regard is glycerol (or otherpolyols), as it is unlikely to denature the enzyme when employed at thehigh concentration to substantially shift the pH min.

8. Optimization Of On and Off Exchange Times

Each peptide amide hydrogen associated with the protein-binding partnerinteraction surface has a unique exchange rate with solvent tritium inthe native folded, unliganded state, which is then shifted to anotherdistinct exchange rate once protein-binding partner complex formationhas occurred. The signal to noise ratio (ratio of tritium functionallybound to this peptide amide over total background tritium bound to allother peptide amides in the protein) can be optimized by a knowledge ofthe exchange rates of this amide hydrogen in the native unligandedprotein and in the protein-binding partner complex.

An amide hydrogen with an exchange half-life of one minute in theprotein's native, unliganded state and 10 minutes in the liganded statemight be optimally studied by on-exchanging the receptor protein for 2minutes (2 half-lives of on-exchange time will result in incorporationof tritium at 75% of the maximal possible equilibrium labeling of thepeptide amide) followed by 10 minutes of off-exchange in the ligandedstate (50% of on-exchanged label will remain on the functionally labeledpeptide amide and less than 0.1% of on-exchanged label will remain oneach of the background labeled peptide amides).

To measure the exchange rates of a particular functionally labelablepeptide amide as it exists in the native, unliganded protein, aliquotsof protein are on-exchanged for varying times (0.5 seconds to 24 hours),bound to binding partner, and then off-exchanged for a fixed time,preferably 24 hours. After pH 2.7, 0° C. proteolytic digestion and HPLCseparation, radioactivity associated with the peptide fragmentcontaining the peptide amide under study is measured. The amount of theradioactivity which represents background (amides which are notfunctionally labeled) is determined by measuring the amount of labelassociated with the same peptide when the protein is on-exchanged forthe same duration but off-exchanged for 24 hours in the absence of addedligand prior to proteolysis and HPLC analysis. Specific radioactivityassociated with the amide is determined as a function of on-exchangetime, and the half-life of (on) exchange of the amide in the unligandedprotein calculated.

To determine the exchange rate of the same peptide amide when it is inthe protein-binding partner complex, protein is on-exchanged for afixed, long period of time (preferably 24 hrs) complexed with bindingpartner, off-exchanged for varying times (preferably 10 seconds to 4days), acid proteolysed, and HPLC analyzed as above. Specificradioactivity associated with the amide is determined as a function ofoff-exchange time, and the half-life of (off)-exchange of the amide inthe liganded protein calculated. With this information the times of onand off-exchange are adjusted to optimize the signal/noise ratio foreach of the amides functionally labeled in the protein-binding partnersystem under study.

9. Modeling of Receptor-Ligand Contact Surfaces

Studies identical in design to those described above (1-8) may also beperformed on the corresponding binding partner protein (the bindingpartner protein is on-exchanged, liganded to receptor protein,off-exchanged, etc.), resulting in the identification of the amides ofthe binding partner which are slowed in exchange by virtue ofinteraction with receptor protein. The knowledge of the identity of theprecise contact peptides in both protein and binding partner may be usedto produce computer-assisted models for the complementary 3-dimensionalstructures of the protein and binding partner surfaces.

Construction of these models is aided by additional information providedby the invention which allows the identification of a subset of peptideamides on the protein's binding surface which are likely to formhydrogen bonds with acceptor residues on the cognate binding proteincontact surface. While most of the peptide amides present on the native,uncomplexed protein or binding partner interaction surfaces can beexpected to be hydrogen bonded to other portions of the same protein, afraction of these peptide amides, possibly approaching 50%, may behydrogen bonded only to solvent. As most protein-binding partner contactsurfaces are highly complementary to each other, it is likely that uponcomplex formation solvent water is removed from the interactionsurfaces, and amides previously hydrogen bonded to water will form newhydrogen bonds to the complementary surface of the partner. This subsetof binding surface amides is readily identified in our studies (Step 8)as they will have an exchange rate in the protein's native, unligandedstate of 0.5 seconds at pH 7.0 and 0° C. These amides can form hydrogenbonds with the complementary surface only if their hydrogens areoriented in the direction of the complementary surface. This in turnplaces orientation constraints on the entire associated peptide bond andto a lesser degree the side chains of the two flanking amino acidresidues of each such amide. Application of these constraints to theforegoing models of interaction surface structure allow higherresolution modeling of the 3-dimensional structure of theprotein-binding partner ligand interaction surfaces.

10. Automation of the Procedures Required for the Performance ofEnzymatic Degradation and HPLC Analysis Under Slowed Tritium ExchangeConditions

While digestion and analysis procedures are performed at 0° C.,analytical samples of tritium exchange-labeled peptides must be storedat temperatures of approximately −60 to −80° C. if unacceptable lossesof label from the peptide are to be avoided over intervals of hours toweeks. Tritium exchange continues in frozen samples in a mannerinversely related to temperature but effectively stops at temperaturesof approximately −70° C. At present, tritium exchange analysis isperformed by manually removing samples from −70° C. storage, meltingthem manually at 0° C., manual addition of reagents (buffers, enzymes)and manual injection of samples onto the HPLC column. Thesemanipulations are labor intensive and expose the samples to inadvertentheating during handling. If HPLC-separated peptides are to be collectedand stored for future study, they are manually collected and stored at−70° C. No presently available robotic HPLC autosampler has thecapability of performing the necessary manipulations on samples storedin the frozen state.

A Spectraphysics AS3000® autosampler may be modified so as to allowautomation of these steps. These preferred modifications were: inclusionof a solid dry ice bath in which samples are stored until analysis; useof modified fluidic syringes which operate reliably at 0° C.; control ofthe autosampler by an external computer; and placement of theautosampler HPLC column and spectrophotometer within a 0° C.refrigerator. Under computerized control, the autosampler's mechanicalarm lifts the desired sample from the −70° C. bath, and places it in aheater/mixer which rapidly melts the sample at 0° C. The liquifiedsample is then automatically injected onto the HPLC column. Operation ofHPLC pumps, on-line radiation counter and data acquisition is similarlyautomated.

To collect tritium-labeled, HPLC-separated peptides under slowedexchange conditions, a Gilson-303® fraction collector (also present inthe 0° C. refrigerator) has been modified so that the sample collectiontubes are immersed in a dry ice bath. Computer-directed diversion ofdesired HPLC effluent fractions into these prechilled tubes results inrapid freezing of the desired tritium-labeled peptides to −70° C.

Deuterium Exchange Embodiments

In another embodiment, functionally labeled proteolytic fragments,generated from a protein that has been functionally labeled withdeuterium (rather than tritium) prior to receptor-ligand complexformation, are analyzed by mass spectroscopy, conducted under conditionswhich minimize off-exchange of peptide amide deuterium from peptidefragments and allow the direct determination of the location offunctionally attached label within a peptide in the size range 3-30amino acids.

Mass spectroscopy has become a standard technology by which the aminoacid sequence of proteolytically generated peptides can be rapidlydetermined (43). It is commonly used to study peptides which containamino acids which have been deuterated at carbon-hydrogen positions, andthereby determine the precise location of the deuterated amino acidwithin the peptide's primary sequence. This is possible because massspectroscopic techniques can detect the slight increase in a particularamino acid's molecular weight due to the heavier mass of deuterium.McCloskey, et al (44) discloses use of deuterium exchange of proteins tostudy conformational changes by mass spectrometry.

The applicant has devised a deuterium-exchange technique essentiallyidentical, in steps 1-5, to the tritium exchange technique describedabove except that on-exchange is performed in deuterated water(preferably 80-99% mole fraction deuterated water). This modifiedprocedure, after addition of binding partner and off-exchange,specifically labels with exchanged deuterium the peptide amides whichmake up the interaction surface between protein and binding partner.Proteolytically generated fragments of protein functionally labeled withdeuterium are identified, isolated, and then subjected to massspectroscopy under conditions in which the deuterium remains in place onthe functionally labeled peptide amides. Standard peptide sequenceanalysis mass spectroscopy can be performed under conditions whichminimize peptide amide proton exchange: samples can be maintained at 4°C. to zero degrees C with the use of a refrigerated sample introductionprobe; samples can be introduced in buffers which range in pH between 1and 3; and analyses are completed in a matter of minutes. MS ions may bemade by MALDI (matrix-assisted laser desorption ionization)electrospray, fast atom bombardment (FAB), etc. The carboxypeptidase mayact before or simultaneously with the ionization events. Subfragmentsare separated by mass by, e.g., magnetic sector, quadropole, ioncyclotron, or time-of-flight methods. For MS methods generally, seeSiuzdak, G., Mass Spectrometry for Biotechnology (Academic Press 1996).

Since deuterium is not radioactive, the deuterium-labeled peptides mustbe identified by other means, such as mass spectrometry (their molecularweight will be greater than that of predicted for the same peptidewithout such a label).

If desired, the same binding protein: binding partner complex may bestudied both by tritium exchange (which need only be to mediumresolution) and by deuterium exchange. The tritium exchange method willidentify the relevant fragments. Since the HPLC mobilities of thesetritium-labeled fragments will then be known, the correspondingdeuterium-labeled fragments can be identified by their common mobilitiesand then subfragmented, etc.

In a preferred embodiment, separate tritium and deuterium exchange runsare avoided. Instead, the deuterated water is supplemented withtritiated water, e.g. the solvent is 98% mole fraction deuterated waterand 2% mole fraction tritiated water (e.g., 50 Ci/ml). As a result, thefragments are labeled both with deuterium and tritium, and the relevantfragments identified by their tritium-imparted radioactivity. Thesubfragments are still analyzed by mass spectroscopy for the presence ofdeuterated label (with appropriate correction for the relatively smallamount of tritium also present). The purpose of the tritium is toradioactively tag peptide fragments containing binding surface residues.However, the exact residues involved are identified by MS analysis ofdeuterium bearing peptides that have been further digested withacid-reactive carboxypeptidases, allowing identification of thedeuterated residues of the radioactive peptides.

In a preferred embodiment, receptor-binding partner complexesfunctionally labeled with deuterium and tritium at their interactionsurface are (under slowed exchanged conditions as described above forhigh resolution tritium exchange analysis) pepsin digested, subjected torpHPLC in 0.1% TFA-containing buffers and column effluent containingtritium labeled peptides subjected to mass spectroscopic analysis. Tomore precisely localize the deuterium label within each peptide, massspectrometry is performed on labeled-proteolytic fragments, that areprogressively further digested (under slowed exchange conditions) withacid reactive carboxypeptidases (41). This digestion can be performedbefore introduction of the sample into the mass spectrometer, orcontinuously in situ while the sample is held in the mass spectrometer.As digestion proceeds, molecular ions of each of the resultingenzyme-generated carboxy-terminal truncated peptide subfragments isdetected by the mass spectrometer, and its molecular weight compared tothat known for the undeuterated form of the same peptide fragment.Peptide fragments which bear functionally attached deuterium areidentified by an increase in their molecular weight of one atomic unitwhen compared to the same peptide fragment generated from undeuteratedreceptor-binding partner. Sufficient subfragmentation and analysis asabove results in the deduction of the protease-generated fragments thathave functionally-bound deuterium. Thereby, the location of eachdeuterated amide within the peptide is determined.

In vivo Analysis

In situ analysis of protein-binding partner interactions is possible invivo. The protein, while present in its native environment as acomponent of an intact living cell, or as a component of a cellularsecretion such as blood plasma, is on-exchanged by incubating cells orplasma in physiologic buffers supplemented with tritiated (ordeuterated) water. The binding partner is then added, allowed to complexto the cell or plasma-associated protein, and then off-exchangeinitiated by returning the cell or plasma to physiologic conditions freeof tritiated (or deuterated) water. During the off-exchange period(hours to days) the formed protein-binding partner complex is isolatedfrom the cell or plasma by any purification procedure which allows thecomplex to remain continuously intact. At the end of the appropriateoff-exchange period, fragmentation and analysis of purified complexproceeds as above.

This analytic method is especially appropriate for proteins which losesubstantial activity as a result of purification, as the binding site islabeled prior to purification.

Binding Site Analysis by Indirect Hydrogen Exchange

In the methods described above, the entire surface of the protein islabeled initially, and label is then removed from those surfaces whichremain solvent exposed after formation of the complex of the bindingprotein and its binding partner. The binding site of the protein isoccluded by the binding partner, and label is therefore retained at thissite.

When the complex is formed, the binding protein may undergo changes inconformation (allosteric changes) at other sites, too. If these changesresult in segments of the protein being buried which, previously, wereon the surface, those segments will likewise retain label.

It is possible to distinguish binding site residues from residuesprotected from “off-exchange” by allosteric effects. In essence, thebinding partner, rather than the binding protein, is labeled initially.The binding protein is labeled indirectly as a result of transfer oflabel from the binding partner to the binding protein. Such transferwill occur principally at the binding surface.

This procedure will functionally label receptor protein amides if theyare slowed by complex formation and are also in intimate contact withthe binding partner in the complexed state. Receptor protein amides thatare slowed because of complex formation-induced allosteric changes inregions of the protein which are not near the protein-binding partnerinteraction surface will not be labeled. This procedure may be performedas follows:

1) binding partner is added to tritiated water (preferably of highspecific activity) to initiate tritium exchange labeling of the bindingpartner.

2) After sufficient labeling is achieved, binding partner is separatedfrom the excess of solvent tritium under conditions which produceminimal loss of tritium label from the binding partner. This can beaccomplished by, e.g., a) shifting the buffer conditions to those ofslowed exchange (0° C., acidic pH) followed by G-25 spin columnseparation of the binding partner into tritium-free buffer or b)employing stopped-flow techniques in which the on-exchange mixture israpidly diluted with large volumes of tritium free buffer.

3) the tritium-labeled binding partner, now essentially free of excesssolvent tritium, is added to receptor protein and conditions adjusted toallow spontaneous reversible (equilibrium) complex formation to takeplace between the two. The conditions of temperature and pH should alsoallow, and preferably maximize, the specific transfer of tritium labelfrom the labelled binding partner to amides on the binding protein'sinteraction surface with partner. Typically, the pH will be 5-8(conducive to ligand binding) and the temperature 0-37° C. Initially,use of pH 7 and 22° C. is recommended, the transfer being controlled bycontrolling the incubation time. A typical trial incubation time wouldbe 24 hours. These conditions of pH, temperature and incubation time mayof course be varied.

4) The complex is then incubated for periods of time sufficient to allowtransfer of tritium label from the labeled binding partner to thereceptor protein. During this incubation period, tritium which hason-exchanged to regions of the binding partner that are distant from thereceptor-binding partner interaction surface will leave the bindingpartner by exchange with solvent hydrogen and be rapidly and highlydiluted in the large volume of solvent water, thereby preventing itsefficient subsequent interaction with the binding protein. However,tritium label that has been attached to binding partner amides presentwithin the (newly formed) protein-binding partner interaction surfacewill be capable of exchanging off of the binding partner only during thebrief intervals when the interaction surface is exposed to solventwater, i.e., when the complex is temporarily dissociated. When sodissociated and solvent exposed, a portion of tritium present on amideswithin the binding partner's interaction surface will leave the surfaceand for a brief time, remain within the proximity of the surface. Giventhe rapid (essentially diffusion limited) rebinding of binding proteinand partner, much of the released tritium that (briefly) remains withinthe environs of the partner's binding surface will in part exchange withamides on the (future) interaction surface of the approaching bindingprotein molecule that subsequently binds to the binding partner. Oncesuch binding occurs, the transferred tritium is again protected fromexchange with solvent until the complex dissociates again. The resultwill be the progressive transfer of a portion of the tritium from thebinding partner interaction surface to exchangeable amides on thecognate protein interaction surface.

Amides whose exchange rates are conformationally slowed each timecomplex formation occurs can also become labelled with tritium, but theywill do so at a much slower rate than amides within the binding surface,as they are located more distant from the high concentration of tritium“released” at the interaction surface with each complex dissociationevent. The efficiency of transfer is roughly inversely proportional tothe cube of the distance between such conformational changes and thebinding surface.

The binding protein-tritiated binding partner complex incubationconditions are adjusted to optimize specific interaction surface amidetritium transfer (SISATT) for a particular binding protein-partner pair.SISATT is defined as the ratio of the amount of tritium (CPM)transferred from binding partner to binding protein peptide amidespreviously determined (by the technique of claim 1) to undergo slowingof amide hydrogen exchange upon binding-protein partner complexformation divided by the total tritium (CPM) transferred from bindingpartner to all peptide amides in the binding protein.

5) After an incubation period that allows and preferably maximizesSISATT, the conditions of slow hydrogen exchange are restored, thecomplex is dissociated and the binding protein fragmented. Fragments ofbinding protein (as opposed to the initially labeled binding partner)that bear tritium label are identified, and further characterized aspreviously described. Alternatively, deuterium is used instead oftritium as the label. Deuterium has the advantage of allowing a muchhigher loading of label (since deuterium is much cheaper than tritium).

It is possible, also, to directly label the binding partner withdeuterium and the binding protein with tritium. As a result, both thebinding site and allosterically buried amides of the binding proteinwill be tritiated, but only binding site amides will be deuterated.

The indirect method is especially applicable to study of proteins whichundergo substantial conformation of changes after, or in the course ofbinding, such as insulin and its receptor.

Compositions

After determining the binding sites of a binding protein or a bindingpartner, by the present methods (alone or in conjunction with othermethods), the information may be exploited in the design of newdiagnostic or therapeutic agents. Such agents may be fragmentscorresponding essentially to said binding sites (with suitable linkersto hold them in the proper spatial relationship if the binding site isdiscontinuous), or to peptidyl or non-peptidyl analogues thereof withthe similar or improved binding properties. Or they may be moleculesdesigned to bind to said binding sites, which may, if desired,correspond to the paratope of the binding partner.

The diagnostic agents may further comprise a suitable label or support.The therapeutic agents may further comprise a carrier that enhancesdelivery or other improves the therapeutic effect.

The agents may present one or more epitopes, which may be the same ordifferent, and which may correspond to epitopes of the same or differentbinding proteins or binding partners.

EXAMPLE

As a demonstration of the practical use of this technology, Applicanthas studied the interaction of human hemoglobin with two differentmonoclonal antibodies known to be reactive with defined and previouslyidentified subregions of the hemoglobin binding protein haptoglobin.These studies employed monoclonal antibody β6-1-23456 (specific for thehuman hemoglobin β chain; epitope centered on or about β6-Glu andmonoclonal antibody β-121 (specific for the human hemoglobin β chain inthe region of residue β-121), both antibodies being the generous gift ofC. R. Kiefer, Medical College of Georgia, Augusta, Ga. (51). Humanhaptoglobin was obtained from Calbiochem Corporation, La Jolla, Calif.

Preparation of hemoglobin: Blood was drawn from a normal donor intosodium heparin at 10 U/ml. Red blood cells were washed five times incold phosphate buffered saline (PBS) (pH 7.4) with the buffy coataspirated after each wash. An equal volume cold distilled water wasadded to the washed cell pellet to lyse cells, and then a one-halfvolume of cold toluene was added with vigorous vortexing. This mixturewas centrifuged for 30 minutes in a cold Sorvall® centrifuge (Dupont)rotor at 15,000 rpm (33,000 g). The hemoglobin (middle) layer wasremoved and the centrifugation and hemoglobin decantation repeated. Theisolated hemoglobin was dialyzed against four changes of cold 0.1Msodium phosphate, 0.5% NaCl pH 7.4. After dialysis, the sample wastreated with carbon monoxide for 15 minutes. Final hemoglobinconcentration was measured by using a molar extinction for heme at 540nm of 14,270. The preparation was stored frozen in aliquots at −70° C.

Preparation of pepsin: Porcine pepsin (Worthington Biochemical Corp.)was dissolved at 10 mg/ml in 50 mM sodium acetate pH 4.5 and dialyzedagainst the same solution to remove proteolytic fragments. It was storedfrozen in aliquots at −70° C.

Tritium exchange: All steps were performed at 0° C. On-exchange wasinitiated by mixing equal volumes (5 μl) of isolated hemoglobin (300mg/ml) and tritiated water (50 Ci/ml) and the mixture incubated for fourhours. Aliquots of this mixture (1.3 μl) were then added to equimolarquantities of either monoclonal β6, monoclonal β121, haptoglobin, (allat 10 mg/ml in PBS, pH 7.4, in a final incubation volume of 75 μl) oradded to 75 μl of PBS alone. These hemoglobin-ligand mixtures were thenimmediately applied to 2 ml Sephadex® G-25 spin columns and centrifuged4 minutes at 1100 g. Spin columns were prepared by filling 3 mlpolypropylene columns (Fisher Scientific) with 2 ml of Sephadex G-25fine equilibrated in PBS pH 7.4 plus 0.1% Triton® X-100. Columns werepre-spun at 1100 g for 2 minutes just before use. After columnseparation, samples were off-exchanged by incubation for a period of 40hours, ten times the length of on-exchange. Samples were then hydrolyzedwith pepsin. Typically, 25 μl of off-exchanged mixture containing 70 μgof hemoglobin was added to 10 μg pepsin in 110 μl of 0.1M NaPO₄ pH 2.7plus 2.5 μl 0.5M H₃PO₄, the mixture incubated on ice for 10 minutes andthen injected onto the HPLC column. An aliquot of on-exchangedhemoglobin was immediately adjusted to pH 2.7, passed over a pH 2.7 (0.1M NaPO₄ pH 2.7) also proteolyzed and analyzed as above without a periodof off-exchange. To measure on-exchange rates of specifically labeledamide protons, hemoglobin was on-exchanged as above but with timeintervals ranging from 10 sec.-18 hours, reacted with ligand, andoff-exchanged for 18 hours. Samples were then proteolyzed, subjected toHPLC as below, and specific label on peptides quantified as a functionof on-exchange time.

High pressure liquid chromatography: Digested samples were analyzed on aWaters HPLC unit modified by putting the column and injector undermelting ice. Mobile phase was prepared using Barnstead nanopure water,Aldrich ultrapure sodium phosphate, J. T. Baker Ultrex® grade HCL andHPLC grade acetonitrile from Burdick & Jackson. Mobile phase consistedof 50 mM NaPO₄ pH 2.7 (solvent A) and a mixture of 20% 50 mM NaPO₄ and80% acetonitrile (ACN) final pH 2.7 (solvent B). Separation of peptideswas achieved using a 30 cm Phenomenex Bondclone® 10 C18 column. Thegradient program started at 100% A 0% B and altered the client to 83% A,17% B over 3.4 minutes. From 3.4 to 6.7 minutes the system ran at aconstant 83% A, 17% B and from 6.7 to 73.3 minutes the programimplemented a linear increase in % B from 17% to 51%. Absorbance wasmonitored at 214 nm with a Waters model 441 detector.

For second dimension separation, peptide peaks bearing specific labelisolated as were collected at 0° C., stored frozen at −70° C., thawed at0° C., mixed with an equal volume of 100 mM PO₄ pH 2.7, and subjected toHPLC as above, except that buffer A was 0.115% trifluoracetic acid (TFA)in H₂O and buffer B was 80% ACN, 20% H₂O, 0.1% TFA. Peaks bearingspecific radiolabel were identified and isolated.

Sample collection: HPLC effluent was collected at the HPLC detectoroutflow with a Gilson model 203® fraction collector. Samples (100 to 400fractions per run) were collected and radioactivity measured by addingfive volumes of Aquamix (ICN Radiochemicals) followed by scintillationcounting. In other studies, on-line liquid scintillation counting wasperformed using a B-RAM flow radiation detector (INUS Inc.).

Peptide identification: HPLC-isolated peptide were analyzed by both gasphase Edman sequencing and amino acid analysis at the UCSD proteinsequencing facility.

Results

Hemoglobin-monoclonal Antibody Epitope Mapping

Hemoglobin was on-exchanged for 4 hours and then either proteolyzedwithout a period of off exchange (FIG. 1b), mixed with equimolarquantity of β6 monoclonal and then off-exchanged for 40 hours (FIG. 1c),mixed with monoclonal β-121 and off-exchanged for 40 hours (data notshown) or off-exchanged 40 hours in the absence of added antibody (FIG.1d). When labeled hemoglobin is examined without a period of offexchange (FIG. 1b), at least 17 radiolabeled peaks were resolved, whichgenerally corresponded to the peaks seen in the optical density trace ofthe same HPLC run (FIG. 1a). When labeled hemoglobin was allowed tofully off exchange without the presence of a protecting monoclonalantibody, all radiolabeled peaks disappeared (FIG. 1d). However, whenlabeled hemoglobin was off-exchanged in the presence of the β6monoclonal, a single unique peak bearing radiolabel was seen indicatingthat this fraction contains the β6 monoclonal antigenic epitope (FIG.1c).

When this peak was subjected to second dimension HPLC in TFA-containingsolvents under slowed proton exchange conditions, two peptides wereresolved by optical density at 214 nm, with only one of these bearingradiolabel (see FIG. 2). This label-bearing peptide was found by gasphase microsequencing and amino acid analysis to represent residues 1-14of the hemoglobin beta chain. Measurement of on-exchange rates oflabeled amides in this peptide demonstrated two rate classes, both ofequal size; one which exchanged on with a half life of less than 10seconds, and another with a half life of approximately 1 hour. Specificactivity measurements indicate that 4.3 amide protons within this 14-merpeptide are slowed by interaction of the β6 antibody with hemoglobin. Asynthetic peptide identical to residues 1-14 of the hemoglobin β chain(β1-14) was synthesized, tritium labeled by proton exchange, andsubjected to graded digestion with carboxypeptidase-P (see FIGS. 6a-j).

Similar studies were performed with hemoglobin off-exchanged afterinteraction with 0-121 monoclonal (FIGS. 3a-c). Three pepsin-generatedpeptides were found to bear tritium label (FIG. 3b). After seconddimension HPLC separation in TFA-containing solvents these peaks weresimilarly resolved from contaminants, sequenced, and found to behemoglobin polypeptides 1-14, β113-128, and β15-31. In preliminaryproton counting studies, approximately two β121 monoclonal-slowedprotons are present in each of these three peptides.

The position of these peptidic regions in the folded hemoglobin tetramerare shown in FIGS. 5a and 5 b. The β6 monoclonal labels six amide bondswhich are present on an externally disposed segment of the foldedhemoglobin molecule (β chain amino acids 1-14) which includes thepreviously characterized target epitope of this monoclonal (β6-9) (51).The β-121 monoclonal labels a total of approximately six protons which,though present on the non-contiguous regions of the linear amino acidsequence of hemoglobin are seen to be surface disposed and located inclose proximity to each other in the folded hemoglobin molecule, andinclude the hemoglobin β chain 121 residue.

Mapping of Hemoglobin-haptoglobin Interaction Sites

When hemoglobin binds to haptoglobin it is known that the hemoglobinmolecule contacts haptoglobin through three non-contiguous peptidicregions which consist of hemoglobin a chain 121-127, β11-25 and β131-146(52,53). We therefore anticipated that pepsin cleavage of hemoglobinlabeled at haptoglobin interaction sites would display between 2 and 10radiolabeled peptides. We therefore performed our haptoglobin studies ata higher level of resolution, accomplished by collection of a largernumber of HPLC fractions (see FIGS. 4a-d). Under these conditions,labeled hemoglobin analyzed without a period of off exchangedemonstrates greater than 33 discernable radiolabeled peaks (FIG. 4b),which again correspond to the optical density tracing (FIG. 4a). Labeledhemoglobin off-exchanged in the presence of haptoglobin produces 7specifically radiolabeled peaks (FIG. 4c) which are not present ifhemoglobin is off-exchanged in the absence of haptoglobin (FIG. 4d).These results indicate that this technology works well with areceptor-like ligand interaction system as complex as that of hemoglobinwith haptoglobin.

Solvent Effect

Synthetic hemoglobin β1-14 peptide was tritium-labeled at all peptideamides by proton exchange, and aliquots of labeled peptide subjected to0° C. HPLC analysis as in FIGS. 1a-d except that a range of solvent pH'swere utilized as indicated below. The percent of original peptide-boundtritium that remained bound to the peptide under each HPLC condition wasthen determined.

pH A solvent B solvent 2.1 0.115% TFA in water 80% ACN, 20% H₂O, 0.1%TFA 2.7 50 mM PO₄, pH 2.7 80% ACN, 20% 50 mM PO₄, pH 2.7 3.5 50 mM PO₄,pH 3.5 80% ACN, 20% 50 mM PO₄, pH 3.5 4.0 50 mM PO₄, pH 4.0 80% ACN, 20%50 mM PO₄, pH 4.0 Tritium retention was about 57% for TFA (pH 2.1), 46%for PO₄ (pH 2.7), 34% for PO₄ (pH 3.5), and 14% for PO₄ (pH 4.0).

The invention is not to be limited in scope by the specific embodimentsdescribed which are intended as single illustrations of individualaspects of the invention, and functionally equivalent methods andcomponents are within the scope of the invention. Indeed, variousmodifications of the invention, in addition to those shown and describedherein, will become apparent to those skilled in the art from theforegoing description and accompanying drawings. Such modifications areintended to fall within the scope of the appended claims.

All references cited herein are incorporated in their entireties byreference for all purposes.

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What is claimed is:
 1. A method of characterizing a binding site of abinding protein of known or determinable amino acid sequence, saidmethod comprising the steps of: (a) selectively labeling peptide amidegroups in a binding site of the binding protein that have beendetermined to be substantially protected from exchange with solventhydrogens in a situation where the binding protein binds its bindingpartner; (b) progressively degrading, under conditions of slow hydrogenexchange, the selectively labeled binding protein to obtain at least oneseries of subfragments, wherein each subfragment of each of said seriesis shorter than the preceding subfragment in the series by about 1-5fewer amino acid residues at one terminus; (c) quantifying an amount ofheavy hydrogen on each subfragment; and (d) correlating the amount oflabel on the subfragments with the amino acid sequence of the bindingprotein, thereby localizing the positions of the binding protein thathad been labeled to within about 1-5 amino acid residues and thuscharacterizing the binding site of the binding protein.
 2. A method ofcharacterizing a binding site of a binding protein of known ordeterminable amino acid sequence, said method comprising the steps of:(a) selectively labeling peptide amide groups in a binding site of thebinding protein so as to obtain a selectively labeled binding proteinwherein either (i) the solvent inaccessible peptide amide groups of thebinding protein comprise a heavy hydrogen and the solvent accessiblepeptide amide groups comprise a normal hydrogen or (ii) the solventinaccessible peptide amide groups of the binding protein comprise anormal hydrogen and the solvent accessible peptide amide groups comprisea heavy hydrogen, and wherein the solvent inaccessible peptide amidegroups have been determined to be those groups that are substantiallyinaccessible to solvent in a situation where the binding protein bindsits binding partner; (b) fragmenting the selectively labeled bindingprotein into a plurality of fragments under conditions of slow hydrogenexchange; (c) determining which fragments are labeled with heavyhydrogen; (d) progressively degrading, under conditions of slow hydrogenexchange, each of the heavy hydrogen-labeled fragments to obtain atleast one series of subfragments, wherein each subfragment of each ofsaid series is shorter than the preceding subfragment in the series byabout 1-5 fewer amino acid residues at one terminus; (e) quantifying anamount of heavy hydrogen on each subfragment; and (f) correlating theamount of heavy hydrogen on the subfragments with the amino acidsequences of the fragments from which the subfragments were generated,thereby localizing the positions of the fragment that had been labeledwith heavy hydrogen to a resolution about 1-5 amino acid residues, andthus characterizing the binding site of the binding protein.
 3. Themethod of claim 1 or 2 wherein the selectively labeling peptide amidegroups of the binding protein comprises the steps of: (i) contacting thebinding protein with a binding partner under conditions wherein thebinding protein binds the binding partner so as to form a bindingprotein-binding partner complex; and (ii) contacting the complex with aheavy hydrogen-labeled solvent under conditions of rapid hydrogenexchange for an “on-exchange” period sufficient for solvent-accessiblepeptide amide hydrogens of the complex to exchange with, and besubstantially replaced by, heavy hydrogens of the solvent, in adetectable number of molecules of the complex.
 4. The method of claim 1or 2 wherein the selectively labeling peptide amide groups of thebinding protein comprises the steps of: (i) contacting the bindingprotein with a heavy hydrogen-labeled solvent under conditions of rapidhydrogen exchange for an “on-exchange” period sufficient forsolvent-accessible peptide amide hydrogens of the binding protein toexchange with, and be substantially replaced by, heavy hydrogens of thesolvent, in a detectable number of molecules of the binding protein;(ii) contacting the binding protein with a binding partner underconditions wherein the heavy hydrogen labels are substantially retainedand wherein the binding protein binds the binding partner so as to forma binding protein-binding partner complex; and (iii) contacting thecomplex with a solvent that is substantially free of heavy hydrogenunder rapid exchange conditions for an “off-exchange” period sufficientfor solvent-accessible peptide amide hydrogens or heavy hydrogens of thecomplex to exchange with, and be substantially replaced by, normalhydrogen of the solvent.
 5. The method of claim 4 wherein the heavyhydrogen is tritium and wherein said quantifying step comprisesradioactivity measurements.
 6. The method of claim 4 wherein the heavyhydrogen is deuterium and wherein said quantifying step comprisesmeasuring the mass of the fragment or subfragment.
 7. The method ofclaim 2, further including the step of separating the fragments prior todetermining which fragments are labeled with heavy hydrogen.
 8. Themethod of claim 7 in which the separation comprises two sequentialseparations steps which are carried out under different conditions. 9.The method of claim 8 in which the first sequential separation step iscarried out at a pH in the range of pH 2.1 to pH 3.0 and the sequentialseparation is carried out at a pH in the range of pH 2.1 to pH 3.0,where the pH of the first and second sequential separation steps aredifferent.
 10. The method of claim 9, in which the first sequentialseparation step is carried out at a pH of 2.7 and the second sequentialseparation step is carried out at a pH of 2.1.
 11. The method of claim 4in which the degradation of the selectively labeled binding protein orlabeled fragments comprises contacting the selectively labeled bindingprotein or labeled fragments with an acid resistant carboxypeptidaseselected from the group consisting of carboxypeptidase P,carboxypeptidase Y, carboxypeptidase W and carboxypeptidase C.
 12. Themethod of claim 4 further comprising disrupting any disulfide bridges inthe selectively labeled binding protein under conditions of slowhydrogen exchange prior to said fragmenting or said progressivelydegrading.
 13. The method of claim 12 wherein said disrupting is carriedout by contacting the selectively labeled binding protein with awater-soluble phosphine at 0-10° C.
 14. The method of claim 4 furthercomprising, prior to fragmenting or progressively degrading, denaturingthe selectively-labeled binding protein under con of slow hydrogenexchange.
 15. The method of claim 14 in which the denaturing is carriedout in a solvent such that the pH for minimization of hydrogen exchangeis substantially higher than that of a purely aqueous solution.
 16. Themethod of claim 15 in which the solvent comprises about 5-20% water andthe remainder is a nonaqueous polar solvent.
 17. The method of claim 16in which the nonaqueous polar solvent is selected from the groupconsisting of acetonitrile, dimethyl sulfoxide, a polyol andcombinations thereof.
 18. The method of claim 17 in which the polyol isglycerol.
 19. The method of claim 16 in which the solvent furtherincludes about 2-4 M guanidine thiocyanate.
 20. The method of claim 16,wherein the pH of the solvent is pH 4.8-5.2.
 21. The method of claim 1or 2 wherein each of said series of subfragments comprises a mixture inwhich each subfragment is present in an analytically sufficient quantityto permit its identification.
 22. The method of claim 1 or 2 whereinsaid progressively degrading is carried out contemporaneously with saidquantifying step.
 23. A method of determining, at a resolution of about1-5 amino acid residues, the position of a peptide amide group labeledwith a heavy hydrogen in a binding site of a binding protein of known ordeterminable amino acid sequence, said method comprising the steps of:(a) selectively labeling peptide amide groups in a binding site of thebinding protein that have been determined to be substantially protectedfrom exchange with solvent hydrogens in a situation where the bindingprotein binds its binding partner; (b) progressively degrading, underconditions of slow hydrogen exchange, said binding protein into amixture of subfragments, wherein said mixture comprises at least oneseries of subfragments, such that each subfragment in each of saidseries is shorter than the preceding subfragment in the series by about1-5 amino acid residues at one terminus and such that the firstsubfragment in the series is derived from and composed of about 1-5fewer amino acid residues than said protein and such that the lastsubfragment in the series is about 1-5 amino acid residues in length;and, when said mixture of subfragments comprises an analyticallysufficient quantity of each subfragment in each of said series, (c)quantifying an amount of heavy hydrogen label on each subfragment; and(d) correlating the amount of heavy hydrogen label of each subfragmentwith the amino acid sequence of the binding protein, thereby localizingthe position of the labeled peptide amide group in the binding site ofthe binding protein to within about 1-5 amino acid residues.
 24. Themethod of claim 1 or 2 wherein said fragmenting is achieved by use of anacid tolerant protease from the Aspergillus protease family.
 25. Themethod of claim 2 wherein the heavy hydrogen label is tritium andwherein said determining comprises radioactivity measurements.
 26. Themethod of claim 2 wherein the heavy hydrogen label is deuterium andwherein said determining comprises measuring the mass of thesubfragment.